Document Outline

)

[1–53]

. The presence of macroscopic

hematuria, persistent or recurrent microscopic hematuria, or a single
episode of microscopic hematuria associated with other abnormal ﬁndings
(eg, stranguria, pollakiuria) is a clear indication for diagnostic testing. A
mild degree of microscopic hematuria (ie, 5–15 red blood cells per high-
power ﬁeld) in the absence of other abnormalities usually is considered to
result from cystocentesis. To exclude iatrogenic hematuria, it is appropriate
to re-evaluate these patients by performing sediment examination on urine
collected during midstream voiding.

Recently, a panel of the American Urological Association was convened to

formulate policy statements for evaluation of asymptomatic microhematuria

E-mail address:

sdforrester@westernu.edu

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.009

Vet Clin Small Anim

34 (2004) 849–866

Box 1. Diseases that may be associated with hematuria in
dogs and cats

Upper urinary tract (kidneys/ureters)

Acute tubular necrosis (aminoglycosides

and ethylene

glycol intoxication

[2,3]

Coagulopathies

Disseminated intravascular coagulation

Coagulation factor deficiencies (hemophilia)
Vitamin K antagonists (coumarin and brodificum)

Thrombocytopenia

[5,6]

Von Willebrand’s disease

Glomerular diseases (acute glomerulonephritis, IgA

glomerulonephropathy, and hereditary nephritis)

[90]

Idiopathic renal hematuria

Leptospirosis

Lyme disease

Nephrolithiasis

Neoplasia (eg, carcinoma, hemangiosarcoma, hemangioma,

sarcoma)

[4,23–27]

Pyelonephritis
Polycystic renal disease
Renovascular abnormalities (renal infarction

and renal

telangiectasia in Welsh Corgi

)

Trauma (blunt force, penetrating wounds, and renal biopsy)

[29–31]

Lower urinary tract (urinary bladder and urethra)

Coagulopathies (upper urinary tract above)

Proliferative urethritis (formerly granulomatous urethritis)

[32,52]

Feline interstitial cystitis

Neoplasia (eg, transitional cell carcinoma,
rhabdomyosarcoma, fibroma)

[32,34–38]

Polypoid cystitis

Urolithiasis

Sterile hemorrhagic cystitis (cyclophosphamide)

[41–43]

Trauma

Diagnostic procedures (urethral catheterization,

cystocentesis, cystoscopy, and voiding
urohydropropulsion)

[44–46]

Ruptured urinary bladder

Urinary tract infection

[47,53]

Vascular ectasia of urinary bladder

[71]

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

in adult human beings

[54–56]

. Microscopic hematuria was deﬁned as three or

more red blood cells per high-power ﬁeld on microscopic evaluation of
urinary sediment from two of three properly collected urine specimens (ie,
freshly voided, clean catch, midstream samples). Human patients with
microscopic hematuria and red blood cell casts, dysmorphic urinary red
blood cells, signiﬁcant proteinuria, or azotemia are evaluated for primary
renal disease

. If there is no evidence of renal disease, a complete urologic

evaluation, including a history, physical examination, laboratory analysis,
voided urinary cytology, radiologic imaging of the upper urinary tract, and
cystoscopy, is recommended

Because some human patients with a negative initial evaluation of

asymptomatic microscopic hematuria eventually develop signiﬁcant uro-
logic disease, follow-up evaluation is recommended

[54]

. For example, the

appearance of hematuria can precede the diagnosis of urinary bladder
cancer by many years in human beings

[57]

. Consideration should be given

to repeating urinalysis and voided urinary cytology at 6, 12, 24, and 36
months

[54]

. Additional evaluation, including repeat diagnostic imaging and

cystoscopy, may be indicated when there is a high index of suspicion for
underlying disease. Immediate urologic re-evaluation is recommended if any
of the following occurs: gross hematuria, abnormal urinary cytology, or
irritative voiding symptoms (eg, dysuria, pollakiuria) in absence of infection.
If none of these occurs within 3 years, additional urologic monitoring is not
considered necessary in human patients

[54]

Studies evaluating the usefulness of diagnostic evaluation of veterinary

patients with subclinical microscopic hematuria are lacking. It seems
reasonable, however, that veterinarians should consider additional evalua-
tion of dogs and cats with persistent microscopic hematuria, even in the
absence of overt clinical signs of urogenital disease. This may be most
appropriate for patients at risk of developing life-threatening disorders, such
as malignant neoplasia (eg, older female dogs, previous treatment with
cyclophosphamide, obesity, exposure to topical ﬂea and tick insecticides)

[58,59]

. Hematuria occurs in 50% to 94% of dogs with urinary bladder

neoplasia and approximately 50% of dogs with renal neoplasia

[25,32,35–

37]

. Although it remains to be studied, it is possible that evaluation of

Urogenital tract (uterus, vagina, vestibule, prostate, and penis)

Estrus
Prostatic disease (eg, benign prostatic hypertrophy, prostatitis,

neoplasia)

[48–50]

Subinvolution of placental sites
Transmissible venereal tumor (vagina, vulva, penis, and

prepuce)

Uterine infection (endometritis and pyometra)
Vaginitis

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

microscopic hematuria could lead to earlier diagnosis and treatment of dogs
and cats with urinary tract neoplasia.

Initial diagnostic evaluation

A diagnostic evaluation of dogs and cats with hematuria is conducted to

localize the site of bleeding within the urogenital tract and to identify the
underlying cause

[60–62]

. This often can be done based on clinical ﬁndings

and results of routine laboratory tests and diagnostic imaging. In some
cases, additional diagnostic testing is needed to determine the source of
bleeding and ﬁnd the underlying cause.

The presence of concomitant signs can be helpful for localizing the source

of hematuria. Pollakiuria, dysuria, or stranguria usually indicates a lower
urinary tract disorder

[32,33,35–37,39,63]

. Infrequently, however, these signs

may be observed in patients with severe renal hematuria that have blood clots
in the urinary bladder and urethra

. Hemorrhagic or serosanguineous

vulvar or preputial discharge may occur in dogs with urethral disease (eg,
proliferative urethritis, neoplasia)

[32,52]

, vaginal or penile neoplasia (eg,

transmissible venereal tumor)

, or prostatic disease

[49,50]

Timing of hematuria

The timing of macroscopic hematuria during urination may help to

localize the cause

[60,61,64]

. Initial hematuria (ie, presence of blood during

the ﬁrst fraction of voided urine) tends to occur with diseases of the urethra
or genital tract, whereas hematuria at the end of urination (ie, terminal
hematuria) suggests a focal lesion in the ventral or ventrolateral aspect of
the urinary bladder (eg, uroliths, polyps). With this type of hematuria, red
blood cells remain in the dependent area of the urinary bladder and are
voided last. In addition to urinary bladder disorders, terminal hematuria
may be observed in patients with intermittent renal hematuria. Hematuria
that occurs throughout urination (ie, total hematuria) may be associated
with coagulopathies or disorders of the kidneys, ureters, or urinary bladder
(especially diﬀuse lesions). It also is possible for total hematuria to occur
secondary to severe disorders of the urethra or prostate gland that cause
reﬂux of blood into the urinary bladder. Lastly, hemorrhagic urethral
discharge may occur independent of urination and be confused with
hematuria in some dogs that have urogenital disease (eg, distal urethral
disease, prostatic disease, vaginal disorders)

[49,64]

Physical examination

A thorough physical examination is indicated in patients with hematuria,

with particular emphasis on signs of coagulopathies and palpation of

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

urogenital organs. Patients with anemia secondary to severe hematuria may
have pale mucous membranes with prolonged capillary reﬁll time. In
addition to hematuria, coagulopathies may cause clinical signs, such as
epistaxis, melena, and prolonged bleeding from venipuncture sites

[65]

Primary hemostatic disorders (eg, thrombocytopenia, von Willebrand’s
disease) may be characterized by petechial or ecchymotic hemorrhages,
whereas coagulation factor deﬁciencies (eg, vitamin K antagonism, hemo-
philia) may also cause hematomas and hemarthrosis

[65,66]

. Abdominal

palpation should focus on examination of the kidneys, urinary bladder,
sublumbar lymph nodes, and prostate gland. A rectal examination should
be performed to evaluate the urethra, caudal urinary bladder, and prostate
gland. In female dogs, a digital vaginal examination may reveal a urethral or
vaginal mass. In males, the prepuce should be retracted fully so that the
mucosal surface of the penis can be thoroughly examined. Male dogs that
are at risk for transmissible venereal tumor (intact, free-roaming, or
outdoors) may have a neoplastic mass located at the base of the penis.
Lastly, the clinician should observe the patient urinate. This may help to
identify or conﬁrm presence of dysuria, stranguria, or pollakiuria as well as
the timing of hematuria during urination. It also allows for collection of
a midstream voided urine specimen, which is indicated in all patients with
hematuria.

Urinalysis

Analysis of urine collected during voiding should be performed initially

to avoid iatrogenic hematuria associated with cystocentesis. If hematuria is
initially identiﬁed in urine collected by cystocentesis, another sample should
be collected during voiding at least 24 hours later. If hematuria is present in
urine collected by free catch but is not identiﬁed in urine obtained by
cystocentesis, diseases of the genital tract (vagina, distal urethra, and penis)
should be suspected. Finding hematuria in urine collected by cystocentesis
as well as in a voided sample indicates bleeding from the kidneys, ureters,
urinary bladder, proximal urethra, or prostate gland. Bleeding from the
latter two sources may reﬂux into the urinary bladder and thus be detected
in urine collected by cystocentesis.

A complete urinalysis should be done, including visual inspection,

biochemical analysis (speciﬁc gravity and dipstick analysis), and microscopic
examination of urine sediment. Initially, urine color and transparency are
evaluated. Patients with macroscopic hematuria have visibly discolored urine
(usually pink, red, reddish brown, or brown) (

). Dipstick analysis

usually reveals a positive reaction for occult blood and protein. The presence
of myoglobin, hemoglobin, and blood in urine may cause a positive occult
blood reaction. Myoglobinuria is rare in dogs and cats but may result from
severe muscle injury (eg, trauma, necrosis, ischemia). Hemoglobinuria may
result from red blood cell lysis in urine (an event that may occur in dilute or

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

alkaline urine) or intravascular hemolysis, when haptoglobin-binding capac-
ity is exceeded. Intravascular hemolysis is associated with pink plasma,
whereas hemoglobinuria secondary to lysis of red blood cells in urine is not.

Although a urine sediment examination may require additional time to

perform, results are helpful and required to conﬁrm the presence of
hematuria. Unless there is local lysis caused by dilute (speciﬁc gravity
1.008) or alkaline urine, red blood cells can be identiﬁed on a urine
sediment examination of patients with hematuria. Depending on the
underlying cause, other abnormalities may be identiﬁed, including pyuria
(urinary tract inﬂammation, infection, or neoplasia), dysplastic or malignant
epithelial cells (urinary tract inﬂammation, infection, or neoplasia), casts
(renal disease or injury), bacteria (urinary tract infection), or crystals
(possible urolithiasis). Red blood cell casts are rarely found on urine
sediment examination; however, their presence indicates bleeding into renal
tubules (ie, renal hematuria) and is probably pathognomonic of glomerular
hemorrhage, a rare cause of hematuria in dogs and cats

[54,64]

. Neoplastic

cells (eg, malignant transitional cells) may be observed on urine sediment
examination in 30% to 70% of dogs with lower urinary tract neoplasia

[35,37]

. The signiﬁcance of these cells must be interpreted cautiously,

because inﬂammation can cause dysplastic changes in epithelial cells,
making them appear malignant

[32,67]

Fig. 1. (A) Urine sample obtained from a 3-month-old female Curly-Coated Retriever with
intermittent macroscopic hematuria. On initial presentation, the urine appeared normal
(transparent and pale yellow). (B) One day later, however, the urine was grossly discolored (dark
reddish brown) and turbid. After extensive diagnostic evaluation, idiopathic renal hematuria was
diagnosed. Right-sided ureteronephrectomy was performed, and the hematuria resolved.

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

Urine culture

Urine culture is indicated in the diagnostic evaluation of dogs and cats with

hematuria. Urinary tract infection (UTI) may cause hematuria, and urinary
tract disorders that cause hematuria (eg, uroliths, neoplasia) may alter host
defense mechanisms and predispose to UTI

[47,53]

. The author’s experience

has been that hematuria is more often microscopic than macroscopic in
patients with UTI. In a study of dogs with UTI associated with hyper-
adrenocorticism or diabetes mellitus, only 2 of 42 dogs (\5%) had grossly
discolored urine reported by the owners, whereas 21 dogs (50%) had
microscopic hematuria

[53]

. If UTI is diagnosed on the basis of urine culture

and hematuria resolves after appropriate antimicrobial treatment, additional
evaluation is probably not needed. If hematuria (either microscopic or
macroscopic) or clinical signs persist despite appropriate treatment of UTI,
however, additional evaluation (eg, diagnostic imaging) should be done.

Urinary bladder antigen test

Recently, a urinary bladder antigen test has been evaluated for the

diagnosis of transitional cell carcinoma (TCC) in dogs

[68–70]

. The test uses

antibodies to detect bladder tumor–associated glycoprotein complex that is
shed in urine of patients with TCC

[68]

. This procedure is sensitive and

speciﬁc for distinguishing between normal dogs and dogs with urinary tract
neoplasia. It is not as reliable for distinguishing between dogs with urinary
tract neoplasia and other urinary tract disorders (eg, UTI, urolithiasis),
however. False-positive results may also occur with signiﬁcant glucosuria
(4

þ), proteinuria (4þ), and pyuria or hematuria (>30–40 white blood cells or

red blood cells per high-power ﬁeld)

[68]

. A negative test result indicates that

TCC is highly unlikely; however, a positive result indicates a need for further
evaluation.

Additional laboratory tests

Depending on suspected causes of hematuria, additional laboratory

testing may be indicated. A complete blood cell count may reveal anemia
in patients with severe hematuria

[71]

. In the author’s experience, severe and

potentially life-threatening anemia secondary to hematuria has most often
occurred in patients with idiopathic renal hematuria, coagulopathies (ie,
severe thrombocytopenia), and renal neoplasia (ie, hemangiosarcoma).
Serum chemistries are indicated to evaluate renal function and detect signs
of underlying diseases (see Box 1).

Coagulation testing is indicated if there is no obvious cause for hematuria

after the initial evaluation (ie, physical examination, routine laboratory
tests, diagnostic imaging) or if there are other signs of bleeding (eg, petechial

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

hemorrhages, epistaxis). If a primary hemostatic defect is suspected,
a platelet estimate or count should be done initially. Hemorrhage associated
with thrombocytopenia generally does not occur unless the platelet count is
less than 50,000 per microliter

[72]

. If the platelet count is normal, a buccal

mucosal bleeding time can be performed to evaluate platelet function and
screen for von Willebrand’s disease. If coagulation factor deﬁciency or
disseminated intravascular coagulation is suspected, coagulation tests,
including prothrombin time (PT), partial thromboplastin time (PTT), ﬁbrin
degradation products (FDPs), and D-dimer concentrations, should be
performed.

Diagnostic imaging

Plain abdominal radiographs and ultrasound are helpful for identifying

and characterizing lesions of the urinary tract that may cause hematuria

[73–76]

. Signs of renal disease (eg, renomegaly, renal masses, nephrolithiasis,

infarcts), prostatic disorders (eg, prostatomegaly), and urocystoliths and
evidence of metastasis (ie, sublumbar lymph node enlargement, vertebral
body or pelvic osteolytic or proliferative lesions) from urogenital neoplasms
may be identiﬁed on abdominal radiographs (

Figs. 2 and 3

). Abdominal

ultrasound may reveal renal architectural changes (

) or urinary

bladder abnormalities (

Figs. 5–7

). Ultrasound may not detect urethral

abnormalities within the pelvic canal; therefore, additional studies, such as
contrast urethrography, may be needed (

Fig. 2. Lateral abdominal radiograph of 12.5-year-old neutered male Poodle with no clinical
signs of urinary disease. Note the presence of several radiopaque urocystoliths. Routine
urinalysis revealed a pH of 8.5, 3

þ proteinuria, 3þ occult blood, 2 to 4 white blood cells per

high-power ﬁeld, 100 to 125 red blood cells per high-power ﬁeld, and malignant transitional
epithelial cells (Courtesy of Johanna Heseltine, DVM, Blacksburg, Virginia).

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

If plain abdominal radiographs or ultrasound fail to demonstrate an

abnormality, contrast radiographic procedures are indicated. If signs of
lower urinary tract disease are present, a double-contrast cystogram and
positive-contrast urethrogram should be done

[73,74]

. Diagnostic imaging

of patients with lower urinary tract neoplasia usually reveals a space-
occupying mass in the urinary bladder or urethra (

Figs. 8 and 9

). Other

abnormalities that may be identiﬁed by contrast radiographic procedures
include uroliths, polypoid cystitis, and blood clots. If renal or ureteral
disease is possible, excretory urography may reveal ﬁndings consistent with
pyelonephritis, hydronephrosis, renal neoplasia, and uroliths.

Fig. 3. Lateral abdominal radiograph of 10-year-old spayed female domestic short-hair cat
with persistent microscopic hematuria. Note the presence of uroliths bilaterally in the renal
pelves. In addition, there appears to be a small urolith in the right proximal ureter.

Fig. 4. Abdominal ultrasound of 7-year-old male Labrador Retriever with a 2.5-week history of
macroscopic hematuria reveals a mass adjacent to the right renal pelvis with some pelvic
enlargement. A right-sided nephrectomy was performed, and histologic evaluation of the renal
mass conﬁrmed hemangiosarcoma.

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

Uroendoscopy

If less invasive tests fail to yield a diagnosis, urethrocystoscopy is indicated

for the evaluation of patients with hematuria, particularly for those with
signs of lower urinary disease (eg, pollakiuria)

[77–81]

. Urethrocystoscopy

allows visualization of the urethral oriﬁce and urethral and urinary bladder
mucosa; collection of tissue samples from masses or other lesions; removal of
uroliths; and collection of urine from individual ureters for diagnosis of renal
hematuria or pyelonephritis (

Fig. 10

). Urethrocystoscopy can be performed

with rigid or ﬂexible cystoscopes in female dogs and cats, whereas only
ﬂexible endoscopes can be used in male animals. The reader is referred
elsewhere for detailed information on urethrocystoscopy

[77–79,81]

Fig. 5. Urinary bladder ultrasound of the 12.5-year-old neutered male Poodle described in

Note the presence of a mass in the trigone (arrows), which was diagnosed as transitional cell
carcinoma. Although not visible in this image, acoustic shadowing of the uroliths also was noted.

Fig. 6. Urinary bladder ultrasound of a 12-year-old neutered male West Highland White Terrier
with a 1-week history of pollakiuria and macroscopic hematuria. Note the presence of
a hyperechoic mass. The mass was surgically removed, and clinical signs resolved. Histopath-
ologic examination of the mass conﬁrmed a leiomyoma, a benign urinary bladder tumor.

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

Cytologic/histologic evaluation of tissue

Collection of tissue for microscopic examination may be needed to

conﬁrm an underlying cause of hematuria in some cases, particularly
neoplasia. Samples for cytologic evaluation may be readily obtained, and

Fig. 7. Urinary bladder ultrasound of a 7-year-old spayed female Miniature Schnauzer with a 2-
day history of macroscopic hematuria, pollakiuria, and stranguria. Note the presence of two
large echogenic densities (mass) within the urinary bladder lumen, presumed to be blood clots.
The physical examination revealed petechial hemorrhages on the ventral abdomen and buccal
mucosa. The platelet count revealed severe thrombocytopenia (\10,000 platelets per
microliter). Results of the diagnostic evaluation were consistent with immune-mediated
thrombocytopenia (IMT). The dog passed numerous blood clots in the urine, and urethral
obstruction occurred on two occasions during hospitalization. The dog responded to treatment
for IMT, and hematuria and echogenic densities in the urinary bladder resolved.

Fig. 8. Contrast urethrogram of a 12-year-old spayed female Miniature Schnauzer with a 2-
month history of pollakiuria, macroscopic hematuria, and hemorrhagic vulvar discharge. The
physical examination, including rectal palpation, revealed no abnormalities. Signiﬁcant
hematuria was not present in urine collected by cystocentesis, and abdominal radiographs
and ultrasound were normal. The urethra appears markedly irregular with multiple ﬁlling
defects throughout its entire length. These ﬁndings were suggestive of either transitional cell
carcinoma or proliferative urethritis.

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

results may yield a diagnosis if numerous criteria of malignancy exist

[67,82]

Concomitant inﬂammation (eg, cystitis), however, may cause dysplastic
changes that mimic neoplasia (eg, malignant transitional cells)

[67]

. The

results of cytologic evaluation should always be interpreted carefully along

Fig. 9. Ventrodorsal radiograph showing the positive-contrast cystogram of a 7-year-old
spayed female Australian Shepherd with a 2-month history of macroscopic hematuria,
pollakiuria, stranguria, weight loss, and inappetence. Note widening of the proximal urethra
(arrows). A urethral mass was identiﬁed during digital rectal palpation, and a necrotic mass was
seen just cranial to the vulva during the physical examination. Histopathologic evaluation of
tissue from the mass revealed ﬁndings consistent with carcinoma.

Fig. 10. (A) Cystoscopy of the dog described in

reveals proliferative and ulcerated tissue

protruding from the urethral oriﬁce. Samples of tissue were collected for cytologic and
histologic evaluation, which conﬁrmed transitional cell carcinoma. (B) For comparison, note
the normal urethral opening seen during vaginoscopy of another patient.

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

with other diagnostic ﬁndings. Histologic examination is preferable when
results of cytologic evaluation are tentative or when more accurate
information would alter treatment or the ability to establish a prognosis
(eg, prognostic information based on tumor grade)

[83]

Tissue specimens for cytologic and histologic evaluation may be collected

by several methods, including percutaneous aspiration, catheter-assisted
biopsy, uroendoscopy, and open biopsy during surgery

[77–81,84–86]

. The

author has most often used ultrasound-guided percutaneous aspiration to
obtain samples for cytologic and histologic evaluation of lesions aﬀecting
the kidneys, urinary bladder, urethra, and prostate gland. Because of the
potential for tumor seeding of the needle tract, however, it has been
recommended that percutaneous aspiration procedures be avoided in
patients with suspected TCC

[59, 87–89]

. An alternative is catheter-assisted

biopsy, which may be useful for patients with mucosal lesions aﬀecting the
urinary bladder, urethra, or prostate gland

[86]

. Ultrasound guidance allows

for more accurate placement of the catheter and may enhance diagnostic
yield

[85]

. The author is not aware of reported occurrences of tumor seeding

along the urethral mucosa; however, this could be a potential complication
of the procedure.

Surgery

If a thorough diagnostic evaluation fails to reveal a cause of hematuria

or if the underlying disease is managed surgically (eg, renal neoplasia),
exploratory celiotomy should be done. Some cases of unilateral renal
hematuria may resolve spontaneously; therefore, another approach would
be to re-evaluate the patient for a speciﬁc period before performing
surgery. If surgery is performed, coagulation abnormalities should be
excluded ﬁrst. Because a nephrectomy may be needed, measurement of
individual kidney glomerular ﬁltration rate (usually by renal scintigraphy)
also should be done before surgery. In the author’s experience, exploratory
surgery is performed when idiopathic renal hematuria is suspected and the
magnitude of hematuria is causing severe clinical signs (eg, weakness
caused by anemia, urinary obstruction secondary to urethral blood clots).
After ventral cystotomy, each ureter is catheterized and urine from each
kidney is evaluated for the presence of blood

. The aﬀected kidney is

removed and submitted for histologic evaluation. If renal hematuria is
not identiﬁed, samples of tissue should be collected from the kidney,
urinary bladder, and prostate gland for histologic evaluation and possibly
bacterial culture, even if they appear normal grossly. If renal hematuria is
bilateral, glomerular disease should be considered and samples of renal
tissue should be collected for light, electron, and immunoﬂuorescence
microscopy (the reader is referred to the article on renal biopsy in this
issue).

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S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

Summary

Hematuria indicates the presence of urogenital disease in dogs and cats.

Persistent hematuria (macroscopic or microscopic) should be evaluated to
determine the source of bleeding and the underlying cause so that
appropriate treatment can be recommended. Results of the history and
physical examination often help to localize disease to the urinary tract
(either upper or lower) or genital tract. Additional diagnostic evaluation,
including laboratory testing (eg, urinalysis, urine culture), diagnostic
imaging (eg, abdominal radiographs, ultrasound), and collection of tissues
for cytologic or histopathologic evaluation, may be needed to identify the
underlying cause. If a thorough evaluation fails to reveal the source or cause
of hematuria, exploratory celiotomy should be considered, especially if
idiopathic renal hematuria is possible.

References

[1] Brown SA, Barsanti JA, Crowell WA. Gentamicin-associated acute renal failure in the

dog. J Am Vet Med Assoc 1985;186(7):686–90.

[2] Grauer GF, Thrall MA, Henre BA, Grauer RM, Hamar DW. Early clinicopathologic

ﬁndings in dogs ingesting ethylene glycol. Am J Vet Res 1984;45(11):2299–303.

[3] Thrall MA, Grauer GF, Mero KN. Clinicopathologic ﬁndings in dogs and cats with

ethylene glycol intoxication. J Am Vet Med Assoc 1984;184(1):37–41.

[4] Crow SE, Bell TG, Wortman JA. Hematuria associated with renal hemangiosarcoma in

a dog. J Am Vet Med Assoc 1980;176(6):531–3.

[5] O’Brien RT, Wood EF. Urinary bladder mural hemorrhage associated with systemic

bleeding disorders in three dogs. Vet Radiol Ultrasound 1998;39(4):354–6.

[6] Garon CL, Scott MA, Selting KA, Cohn LA. Idiopathic thrombocytopenic purpura in

a cat. J Am Anim Hosp Assoc 1999;35(6):464–70.

[7] Batamuzi EK, Kristensen F, Basse A, Dahl S. Idiopathic renal haematuria in a dog.

Vet Rec 1994;135(25):603.

[8] Hitt ME, Straw RC, Lattimer JC, Shaw DP, Weide KD, Smith C. Idiopathic hematuria of

unilateral renal origin in a dog. J Am Vet Med Assoc 1985;187(12):1371–3.

[9] Holt PE. Idiopathic renal haematuria in the dog. Vet Rec 1995;136(2):52.

[10] Jennings PB Jr, Mathey WS, Okerberg CV, Langlinais PC, Kim SH. Idiopathic renal

hematuria in a military working dog. Mil Med 1992;157(10):561–4.

[11] Kaufman AC. Benign essential hematuria in dogs. Compend Contin Educ Pract Vet 1994;

16(10):1317–23.

[12] Hawthorne JC, deHaan JJ, Goring RL, Randall SR, Kennedy FS, Stone E, et al.

Recurrent urethral obstruction secondary to idiopathic renal hematuria in a puppy. J Am
Anim Hosp Assoc 1998;34(6):511–4.

[13] Stone EA, DeNovo RC, Rawlings CA. Massive hematuria of nontraumatic renal origin in

dogs. J Am Vet Med Assoc 1983;183(8):868–71.

[14] Mishina M, Watanabe T, Yugeta N, Maeda H, Fujii K, Wakao Y, et al. Idiopathic renal

hematuria in a dog; the usefulness of a method of partial occlusion of the renal artery.
J Vet Med Sci 1997;59(4):293–5.

[15] Adin CA, Cowgill LD. Treatment and outcome of dogs with leptospirosis: 36 cases (1990–

1998). J Am Vet Med Assoc 2000;216(3):371–5.

862

S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

[16] Birnbaum N, Barr SC, Center SA, Schermerhorn T, Randolph JF, Simpson KW.

Naturally acquired leptospirosis in 36 dogs: serological and clinicopathological features.
J Small Anim Pract 1998;39(5):231–6.

[17] Langston CE, Heuter KJ. Leptospirosis. A re-emerging zoonotic disease. Vet Clin North

Am Small Anim Pract 2003;33(4):791–807.

[18] Rentko VT, Clark N, Ross LA, Schelling SH. Canine leptospirosis. A retrospective study

of 17 cases. J Vet Intern Med 1992;6:235–44.

[19] Littman MP. Canine borreliosis. Vet Clin North Am Small Anim Pract 2003;33(4):827–62.
[20] Grauer GF, Burgess EC, Cooley AJ, Hagee JH. Renal lesions associated with Borrelia

burgdorferi infection in a dog. J Am Vet Med Assoc 1988;193(2):237–9.

[21] Dambach DM, Smith CA, Lewis RM, Van Winkle TJ. Morphologic, immunohistochemical,

and ultrastructural characterization of a distinctive renal lesion in dogs putatively associated
with Borrelia burgdorferi infection: 49 cases (1987–1992). Vet Pathol 1997;34(2):85–96.

[22] DiBartola SP, Myer CW, Boudrieau RJ, DeHoﬀ WD. Chronic hematuria associated with

renal pelvic blood clots in two dogs. J Am Vet Med Assoc 1983;183(10):1102–4.

[23] Eddlestone S, Taboada J, Senior D, Paulsen DB. Renal haemangioma in a dog. J Small

Anim Pract 1999;40(3):132–5.

[24] Henry CJ, Turnquist SE, Smith A, Graham JC, Thamm DH, O’Brien M, et al. Primary

renal tumours in cats: 19 cases (1992–1998). J Feline Med Surg 1999;1(3):165–70.

[25] Klein MK, Cockerell GL, Harris CK, Withrow SJ, Lulich JP. Canine primary renal

neoplasms: a retrospective review of 54 cases. J Am Anim Hosp Assoc 1988;24:443–52.

[26] Seaman RL, Patton CS. Treatment of renal nephroblastoma in an adult dog. J Am Anim

Hosp Assoc 2003;39(1):76–9.

[27] Wang FI, Su HL. A renal hemangiosarcoma causing hematuria in a dog. Proc Natl Sci

Counc Repub China B 2001;25(3):187–92.

[28] Moore FM, Thornton GW. Telangiectasia of Pembroke Welsh Corgi dogs. Vet Pathol

1983;20(2):203–8.

[29] Grauer GF, Twedt DC, Mero KN. Evaluation of laparoscopy for obtaining renal biopsy

specimens from dogs and cats. J Am Vet Med Assoc 1983;183(6):677–9.

[30] Jeraj K, Osborne CA, Stevens JB. Evaluation of renal biopsy in 197 dogs and cats. J Am

Vet Med Assoc 1982;181(4):367–9.

[31] Weisse C, Aronson LR, Drobatz K. Traumatic rupture of the ureter: 10 cases. J Am Anim

Hosp Assoc 2002;38(2):188–92.

[32] Moroﬀ SD, Brown BA, Matthiesen DT, Scott RC. Inﬁltrative urethral disease in female

dogs: 41 cases (1980–1987). J Am Vet Med Assoc 1991;199(2):247–51.

[33] Buﬃngton CA, Chew DJ, Kendall MS, Scrivani PV, Thompson SB, Blaisdell JL, et al.

Clinical evaluation of cats with nonobstructive urinary tract diseases. J Am Vet Med Assoc
1997;210(1):46–50.

[34] Brearley MJ, Thatcher C, Cooper JE. Three cases of transitional cell carcinoma in the cat

and a review of the literature. Vet Rec 1986;118(4):91–4.

[35] Burnie AG, Weaver AD. Urinary bladder neoplasia in the dog: a review of seventy cases.

J Small Anim Pract 1983;24:129–43.

[36] Esplin DG. Urinary bladder ﬁbromas in dogs: 51 cases (1981–1985). J Am Vet Med Assoc

1987;190(4):440–4.

[37] Norris AM, Laing EJ, Valli VE, Withrow SJ, Macy DW, Ogilvie GK, et al. Canine bladder

and urethral tumors: a retrospective study of 115 cases (1980–1985). J Vet Intern Med
1992;6(3):145–53.

[38] Wimberly HC, Lewis RM. Transitional cell carcinoma in the domestic cat. Vet Pathol

1979;16(2):223–8.

[39] Martinez I, Mattoon JS, Eaton KA, Chew DJ, DiBartola SP. Polypoid cystitis in 17 dogs

(1978–2001). J Vet Intern Med 2003;17(4):499–509.

[40] Collins RL, Birchard SJ, Chew DJ, Heuter KJ. Surgical treatment of urate calculi in

Dalmatians: 38 cases (1980–1995). J Am Vet Med Assoc 1998;213(6):833–8.

863

S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

[41] Crow SE, Thielen GH, Madewell BR, Weller RE, Henness AM. Cyclophosphamide-

induced cystitis in the dog and cat. J Am Vet Med Assoc 1977;171(3):259–62.

[42] Peterson JL, Couto CG, Hammer AS. Acute sterile hemorrhagic cystitis after a single

intravenous administration of cyclophosphamide in three dogs. J Am Vet Med Assoc 1992;
201:1572–4.

[43] Laing EJ, Miller CW, Cochrane SM. Treatment of cyclophosphamide-induced hemor-

rhagic cystitis in ﬁve dogs. J Am Vet Med Assoc 1988;193(2):233–6.

[44] Lulich JP, Osborne CA, Carlson M, Unger LK, Samelson LL, Koehler LA, et al.

Nonsurgical removal of urocystoliths in dogs and cats by voiding urohydropropulsion.
J Am Vet Med Assoc 1993;203(5):660–3.

[45] Lulich JP, Osborne CA, Sanderson SL, Ulrich LK, Koehler LA, Bird KA, et al. Voiding

urohydropropulsion. Lessons from 5 years of experience. Vet Clin North Am Small Anim
Pract 1999;29(1):283–91.

[46] Kruger JM, Osborne CA, Ulrich LK. Cystocentesis. Diagnostic and therapeutic

considerations. Vet Clin North Am Small Anim Pract 1996;26(2):353–61.

[47] Seguin MA, Vaden SL, Altier C, Stone E, Levine JF. Persistent urinary tract infections and

reinfections in 100 dogs (1989–1999). J Vet Intern Med 2003;17(5):622–31.

[48] Krawiec DR, Heﬂin D. Study of prostatic disease in dogs: 177 cases (1981–1986). J Am Vet

Med Assoc 1992;200(8):1119–22.

[49] Read RA, Bryden S. Urethral bleeding as a presenting sign of benign prostatic

hyperplasia in the dog: a retrospective study (1979–1993). J Am Anim Hosp Assoc 1995;
31(3):261–7.

[50] Ling GV, Nyland TG, Kennedy PC, Hager DA, Johnson DL. Comparison of two sample

collection methods for quantitative bacteriologic culture of canine prostatic ﬂuid. J Am Vet
Med Assoc 1990;196(9):1479–82.

[51] Rogers KS, Walker MA, Dillon HB. Transmissible venereal tumor: a retrospective study

of 29 cases. J Am Anim Hosp Assoc 1998;34:463–70.

[52] Hostutler RA, Chew DJ, Eaton KA, DiBartola SP. Cystoscopic appearance of

proliferative urethritis in 2 dogs before and after treatment. J Vet Intern Med 2004;
18(1):113–6.

[53] Forrester SD, Troy GC, Dalton MN, Huﬀman JW, Holtzman G. Retrospective evaluation

of urinary tract infection in 42 dogs with hyperadrenocorticism or diabetes mellitus or
both. J Vet Intern Med 1999;13(6):557–60.

[54] Grossfeld GD, Wolf JS, Litwin MS, Hricak H, Shuler CL, Agerter DC, et al.

Asymptomatic microscopic hematuria in adults: summary of the AUA best practice
policy recommendations. Am Fam Physician 2001;63:1145–54.

[55] Grossfeld GD, Litwin MS, Wolf JS Jr, Hricak H, Shuler CL, Agerter DC, et al.

Evaluation of asymptomatic microscopic hematuria in adults: the American Urological
Association best practice policy–part II: patient evaluation, cytology, voided markers,
imaging, cystoscopy, nephrology evaluation, and follow-up. Urology 2001;57(4):
604–10.

[56] Grossfeld GD, Litwin MS, Wolf JS, Hricak H, Shuler CL, Agerter DC, et al. Evaluation of

asymptomatic microscopic hematuria in adults: the American Urological Association best
practice policy–part I: deﬁnition, detection, prevalence, and etiology. Urology 2001;57(4):
599–603.

[57] Hiatt R, Ordonez J. Dipstick urinalysis screening, asymptomatic microhematuria, and

subsequent urological cancers in a population-based sample. Cancer Epidemiol
Biomarkers Prev 1994;3:439–43.

[58] Mutsaers AJ, Widmer WR, Knapp DW. Canine transitional cell carcinoma. J Vet Intern

Med 2003;17(2):136–44.

[59] Henry CJ. Management of transitional cell carcinoma. Vet Clin North Am Small Anim

Pract 2003;33:597–613.

864

S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

[60] Crow SE. Hematuria: an algorithm for diﬀerential diagnosis. Compend Contin Educ Pract

Vet 1980;2(12):211–8.

[61] Grauer GF. Urinary tract disorders. In: Nelson RW, Feldman EC, editors. Small animal

internal medicine. 3rd edition. St. Louis: Mosby; 2003. p. 572–5.

[62] Lage AL. Diagnostic approach to canine and feline hematuria. In: Kirk R, editor.

Current veterinary therapy (small animal practice). vol. X. Philadelphia: WB Saunders;
1989. p. 1117–23.

[63] Kruger JM, Osborne CA, Goyal SM, Wickstrom SL, Johnston GR, Fletcher TF, et al.

Clinical evaluation of cats with lower urinary tract disease. J Am Vet Med Assoc 1991;
199(2):211–6.

[64] Osborne CA, Stevens JB. Urine sediment: under the microscope. In: Osborne CA, Stevens

JB, editors. Urinalysis: a clinical guide to compassionate patient care. Shawnee Mission:
Bayer Corporation; 1999. p. 125–50.

[65] Ruiz de Gopegui R, Feldman BF. Platelets and von Willebrand’s disease. In: Ettinger SJ,

Feldman EC, editors. Textbook of veterinary internal medicine. Philadelphia: WB
Saunders; 2000. p. 1817–28.

[66] Brooks M. Coagulopathies and thrombosis. In: Ettinger SJ, Feldman EC, editors.

Textbook of veterinary internal medicine. Philadelphia: WB Saunders; 2000. p. 1829–41.

[67] Meinkoth JH, Cowell RL. Recognition of basic cell types and criteria of malignancy. Vet

Clin North Am Small Anim Pract 2002;32(6):1209–35.

[68] Borjesson DL, Christopher MM, Ling GV. Detection of canine transitional cell carcinoma

using a bladder tumor antigen urine dipstick test. Vet Clin Pathol 1999;28(1):33–8.

[69] Billet JP, Moore AH, Holt PE. Evaluation of a bladder tumor antigen test for the diagnosis

of lower urinary tract malignancies in dogs. Am J Vet Res 2002;63(3):370–3.

[70] Henry CJ, Tyler JW, McEntee MC, Stokol T, Rogers KS, Chun R, et al. Evaluation of

a bladder tumor antigen test as a screening test for transitional cell carcinoma of the lower
urinary tract in dogs. Am J Vet Res 2003;64(8):1017–20.

[71] Littman MP, Niebauer GW, Hendrick MJ. Macrohematuria and life-threatening anemia

attributable to subepithelial vascular ectasia of the urinary bladder in a dog. J Am Vet Med
Assoc 1990;196(9):1487–9.

[72] Prater R, Tvedten H. Hemostatic abnormalities. In: Willard MD, Tvedten H, editors.

Small animal clinical diagnosis by laboratory methods. Philadelphia: WB Saunders; 2004.
p. 92–112.

[73] Pechman R. The urethra. In: Thrall D, editor. Textbook of veterinary diagnostic

radiology. Philadelphia: WB Saunders; 2002. p. 588–92.

[74] Park R, Wrigley R. The urinary bladder. In: Thrall D, editor. Textbook of veterinary

diagnostic radiology. Philadelphia: WB Saunders; 2002. p. 571–87.

[75] Leveille R. Ultrasonography of urinary bladder disorders. Vet Clin North Am Small Anim

Pract 1998;28(4):799–821.

[76] Feeney D, Johnston G. The kidneys and ureters. In: Thrall D, editor. Textbook of

veterinary diagnostic radiology. Philadelphia: WB Saunders; 2002. p. 556–71.

[77] Brearley MJ, Cooper JE. The diagnosis of bladder disease in dogs by cystoscopy. J Small

Anim Pract 1987;28:75–85.

[78] Cannizzo KL, McLoughlin MA, Chew DJ, DiBartola SP. Uroendoscopy. Evaluation of

the lower urinary tract. Vet Clin North Am Small Anim Pract 2001;31(4):789–807.

[79] Chew DJ, Buﬃngton T, Kendall MS, Osborn SD, Woodsworth BE. Urethroscopy,

cystoscopy, and biopsy of the feline lower urinary tract. Vet Clin North Am Small Anim
Pract 1996;26(3):441–62.

[80] McCarthy TC. Cystoscopy and biopsy of the feline lower urinary tract. Vet Clin North Am

Small Anim Pract 1996;26(3):463–82.

[81] Senior DF. Cystoscopy. In: Tams T, editor. Endoscopy. St. Louis: Mosby; 1999. p. 447–59.
[82] Borjesson DL. Renal cytology. Vet Clin North Am Small Anim Pract 2003;33(1):119–34.

865

S.D. Forrester / Vet Clin Small Anim 34 (2004) 849–866

[83] Valli VE, Norris A, Jacobs RM, Laing E, Withrow S, Macy D, et al. Pathology of canine

bladder and urethral cancer and correlation with tumour progression and survival. J Comp
Pathol 1995;113(2):113–30.

[84] Hager DA, Nyland TG, Fisher P. Ultrasound-guided biopsy of the canine liver, kidney,

and prostate. Vet Radiol 1985;26(3):82–8.

[85] Lamb CR, Trower ND, Gregory SP. Ultrasound-guided catheter biopsy of the lower

urinary tract: technique and results in 12 dogs. J Small Anim Pract 1996;37(9):413–6.

[86] Melhoﬀ T, Osborne C. Catheter biopsy of the urethra, urinary bladder, and prostate gland.

In: Kirk R, editor. Current veterinary therapy, vol. VII. Philadelphia: WB Saunders; 1977.
p. 1173–5.

[87] Anderson WI, Dunham BM, King JM, Scott DW. Presumptive subcutaneous surgical

transplantation of a urinary bladder transitional cell carcinoma in a dog. Cornell Vet 1989;
79(3):263–6.

[88] Gilson SD, Stone EA. Surgically induced tumor seeding in eight dogs and two cats. J Am

Vet Med Assoc 1990;196(11):1811–5.

[89] Nyland TG, Wallack ST, Wisner ER. Needle-tract implantation following ultrasound-

guided ﬁne-needle aspiration biopsy of transitional cell carcinoma of the bladder, urethra,
and prostate. Vet Radiol Ultrasound 2002;43(1):50–3.

[90] Harris CH, Krawiec DR, Gelberg HB, Shapiro SZ. Canine IgA glomerulonephropathy.

Vet Immunol Immunopathol 1993;36(1):1–16.

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Early diagnosis of renal disease and

renal failure

George E. Lees, DVM, MS

Department of Small Animal Medicine and Surgery, College of Veterinary Medicine,

Texas A&M University, 4474 TAMU, College Station, TX 77843–4474, USA

Chronic kidney disease (CKD) is a common problem that causes consider-

able morbidity and mortality in dogs and cats. Historically, veterinarians
often have not detected the presence of ongoing renal diseases in these animals
until the conditions have advanced near their end stages and have caused the
aﬀected animals to develop a degree of renal failure that is suﬃcient to be
manifested by clinical signs of uremia. Treatment of animals that have or are
approaching end-stage renal disease (ESRD) is predictably frustrating,
because therapeutic options generally are limited to palliative or supportive
care and even small additional increments of disease progression, which
usually occur, produce relatively large (as a proportion of residual function)
amounts of further renal impairment and worsening uremia. Successful renal
replacement, such as with dialysis or renal transplantation, eventually is the
only potentially eﬀective treatment for most patients with ESRD. Renal
replacement is unlikely to become a readily available treatment option for
most dogs and cats with CKD, however.

The main goal of early diagnosis of renal disease and renal failure in dogs

and cats is to enable timely application of therapeutic interventions that may
slow or halt disease progression. The fundamental idea is to disconnect the
condition of ‘‘having renal disease’’ from the outcome of ‘‘developing end-
stage renal failure.’’ Absent any potentially beneﬁcial intervention, an early
diagnosis of renal disease or renal failure would merely permit one to observe
a larger portion of the entire disease course. This may have some merit, but the
greater value of early diagnosis lies in its potential to identify animals for
which particular treatments may retard or stop progressive renal damage and
thus preserve adequate renal function for a longer period than would
otherwise be the case. Fortunately, although much remains to be learned,
research in this ﬁeld has begun to arm veterinarians with evidence-based

E-mail address:

glees@cvm.tamu.edu

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.004

Vet Clin Small Anim

34 (2004) 867–885

therapeutic recommendations for combating progression of renal disease in
dogs and cats

[1–9]

. Moreover, one major theme of contemporary research in

nephrology is better deﬁnition of mechanisms underlying renal disease
progression and, thereby, identiﬁcation of additional therapeutic targets
and treatment strategies to be tested. Progress in this arena is likely to have
future application in the care of dogs and cats with CKD.

With a growing armamentarium of established and emerging treatment

strategies to slow progression of CKD in dogs and cats, veterinarians must
become more astute and discriminating detectors of early renal disease or
renal failure if their patients are to beneﬁt optimally from such treatments.
Early diagnosis confers two crucial advantages for treatment eﬃcacy. First,
because there is more residual function left to work with, early intervention
has a much greater impact on the length of survival even if the eﬀect of
treatment on the rate of disease progression is similar whether it is applied
early or late (

). Additionally, most therapeutic strategies are likely to be

more eﬀective (ie, reduce [ﬂatten] the slope of lines representing renal function
over time, as shown in

, to a greater degree) when they are applied early

rather than late in the course of disease. The goal of this article is to review
available methods for early detection and monitoring of renal disease and
renal failure.

General concepts

Existence of renal disease is deﬁned by the presence of lesions in the

parenchyma of one or both kidneys. Most renal lesions arise as a result of

Fig. 1. Hypothetic eﬀects of altering the rate of disease progression at early (point A) versus
later (point B) time in the course of renal disease. In the diagramed example, a renal disease is
rapidly progressing (represented by the steeply sloping line) toward death at time T

. Equally

eﬀective treatments to slow progression (ie, to similarly ﬂatten the slope of the line representing
ongoing renal deterioration) are applied at both time points, but such intervention results in an
approximately fourfold longer prolongation of survival (from time T

to time T

rather than

just to time T

) when instituted early on (at point A) rather than later (point B).

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

injury to or deterioration of one or more of the four major components of
normal kidneys (vessels, glomeruli, tubules, and interstitium), but some renal
lesions are the result of defective renal development. Additionally, most renal
lesions are manifested as changes in the appearance of renal tissue that can be
recognized during gross, light microscopic, or electron microscopic exami-
nations. Some renal lesions exist only at a biochemical or molecular level (eg,
defective transporters in the cell membranes of tubular epithelial cells),
however, and are not manifested by morphologic changes, at least initially.

Although the presence of renal lesions deﬁnes the existence of renal disease,

antemortem discovery of renal disease is uncommonly accomplished by ﬁrst
detecting the renal lesions (eg, structural changes in the kidneys) themselves.
Nonetheless, there are important occasions when this occurs. For example,
abnormal renal size, shape, or ﬁrmness may be detected by palpation during
physical examination. Diagnostic imaging of the abdomen (eg, sonography,
radiology) also may demonstrate evidence of previously unsuspected changes
in renal structure, and these opportunities to discover the existence of renal
disease should not be overlooked. The most common way that renal disease
discovery occurs, however, is by ﬁrst detecting evidence of changes in renal
function that arise as a consequence of renal lesions.

The main physiologic purpose of the kidneys is to regulate the volume and

composition of the extracellular ﬂuid (ECF) compartment to promote
homeostasis. This task is accomplished by glomerular ﬁltration, followed
by tubular reabsorption and secretion. Each of these categories of renal
function encompasses a plethora of speciﬁc individual functional contribu-
tions made by the component cells and extracellular matrices of the kidneys.
Additionally, the kidneys have an important role as endocrine organs, being
sources of hormones (eg, renin, erythropoietin, 1,25-dihydroxycholecalci-
ferol) as well as targets for hormones (eg, aldosterone, antidiuretic hormone).
Many of these component functions of kidneys can become impaired by renal
disease without altering the ability of the kidneys to serve their main function,
that is, to adequately regulate the volume and composition of the ECF.
Reduction of glomerular ﬁltration rate (GFR) by renal disease does lead to an
inability of the kidneys to maintain normal ECF composition, however, and
when this occurs, nitrogenous wastes (eg, urea, creatinine) are the ﬁrst
substances to develop abnormal (ie, increased) concentrations in the ECF.
Therefore, existence of renal failure can be deﬁned by the presence of
azotemia that is caused by renal disease. To identify renal failure, one must
exclude prerenal and postrenal causes of the azotemia; however, ﬁnding
azotemia, especially an increased plasma creatinine concentration, that is
attributable to renal disease indicates that the kidneys are failing to serve their
principal physiologic function adequately.

Fortunately for patients but unfortunately for diagnosticians, normal

kidneys have huge functional reserves. That is, a renal disease can extensively
damage the kidneys without causing evident impairment of renal function.
For example, a healthy individual can relinquish an entire kidney, whether as

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

a renal transplant donor or as a result of its destruction by disease, without
notably altering ordinary indices of renal function. An additional perspective
is gained by considering the amount of destruction and removal of renal
tissue required to achieve ‘‘mild’’ azotemia (plasma creatinine concentrations
about 2–4 mg/dL) in initially healthy dogs (having plasma creatinine con-
centrations about 1 mg/dL) to create the remnant kidney model of CKD in
these animals. To achieve this degree of azotemia after compensatory
hypertrophy of the remaining renal tissue has occurred, investigators have
to destroy 11/12 to 15/16 of one kidney and completely remove the other
kidney

. From this perspective, one can appreciate that initial discovery of

CKD that has already led to development of a plasma creatinine concentra-
tion on the order of 2 to 4 mg/dL despite compensatory changes, which can be
presumed to have been exhausted during the course of a chronic disease
process, does not constitute ‘‘early diagnosis’’ of renal disease. Indeed, this is
a rather late stage in the course of any chronic progressive renal disease
leading to failure.

Potentially eﬀective strategies for early diagnosis of renal disease based on

detection of changes in renal function involve evaluation of urine tests and
blood tests. The urine tests of greatest utility are those that detect (1)
proteinuria that is a manifestation of altered glomerular permselectivity and
(2) impaired urine-concentrating ability. Blood tests of greatest utility are
those that serve as an indirect index of GFR (eg, plasma creatinine con-
centration). These can be supplemented by various methods for estimating
GFR directly; however, all methods for direct estimation of GFR are too
cumbersome and costly for widespread use as screening tests. Using each of
these approaches for early detection of renal disease is discussed sub-
sequently.

Inherent dilemmas

Before considering speciﬁc testing strategies for early diagnosis of renal

disease and renal failure, the dilemmas that are inherent to this task must be
appreciated. First, there is the issue of diagnostic test sensitivity versus
speciﬁcity, and, second, there is the issue of possibly detecting mild non-
progressive renal disease.

Sensitivity versus speciﬁcity of diagnostic tests

In general, the use of ‘‘test X’’ for diagnosis of the presence or absence of

‘‘condition A’’ involves applying a cutoﬀ value to the results of test X to
obtain one of two conclusions: that the test subject does not have condition A
(test result is at or below the cutoﬀ value) or that the test subject does have
condition A (test result is above the cutoﬀ value). For some tests, there is
a single cutoﬀ value that yields excellent diagnostic sensitivity (ie, no false-

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

negative results) and diagnostic speciﬁcity (ie, no false-positive results). This
occurs when there is no overlap in test result values that can be obtained from
individuals that are aﬀected versus unaﬀected by condition A. For many tests,
however, including most of the tests that can be used to screen dogs and cats
for renal disease, the test result values that can be obtained from aﬀected
versus unaﬀected animals do overlap. In this circumstance, the cutoﬀ value
that is used for interpretation of the test results greatly inﬂuences test
sensitivity and speciﬁcity in a reciprocal fashion. Setting the cutoﬀ value near
the low end of the overlapping range reduces false-negative results (ie,
improves sensitivity) but also increases false-positive results (ie, reduces
speciﬁcity). Conversely, choosing a cutoﬀ value near the high end of the
overlapping range reduces false-positive results (ie, improves speciﬁcity) but
also increases false-negative results (ie, reduces sensitivity).

When using a test for deﬁnitive diagnosis, speciﬁcity is paramount, and

choosing a cutoﬀ value that is at or just above the high end of the overlapp-
ing range works best. Historically, this is the diagnostic mode in which
veterinarians most often have been trained to function. Veterinarians seeking
to establish the speciﬁc cause of an animal’s illness appropriately use test
cutoﬀ values that maximize speciﬁcity (ie, minimize false-positive results). In
contrast, for screening groups of animals to detect all those aﬀected by
a particular condition, the superior cutoﬀ values to use are those that
maximize sensitivity (ie, minimize false-negative results). Because an eﬀective
screening strategy often lacks speciﬁcity (ie, identiﬁes some false-positive
results as well as all true-positive results), it must be combined with an
eﬀective follow-up evaluation strategy to identify the animals that actually
are aﬀected by the condition in question. Two general approaches can be used
for such follow-up evaluations. One possibility is to perform an additional
diﬀerent test that is highly speciﬁc for the condition. A second possibility is to
perform serial evaluations of the initial test to detect changes in the observed
results as they rise toward or exceed the cutoﬀ value that has high diagnostic
speciﬁcity.

When performing screening tests, the estimated frequency of the condition

in the group or population of animals being evaluated is yet another
important consideration. Consider two groups of 1000 animals that diﬀer
only in that the prevalence of a condition in the ﬁrst group is 30%, whereas in
the second group, it is merely 3%. Using a screening test that is 100%
sensitive (ie, gives a true-positive result in all aﬀected animals) and 95%
speciﬁc (ie, gives a false-positive result in 5 of 100 unaﬀected animals), there
will be about 335 positive tests in the ﬁrst group, and only 35 of the test results
will be false-positive. In the second group, however, the same screening test
will yield about 78 positive test results, but 48 of these results will be false-
positive. This example illustrates that the importance of a follow-up eval-
uation strategy to diﬀerentiate true-positive from false-positive results is
greater when a screening test is used in groups of animals having a low
prevalence of the condition in question.

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Progressive versus nonprogressive renal disease

If one diagnoses renal disease or renal failure only when the condition is

near end stage, little consideration need be given to the possibility that the
condition might be nonprogressive. There are at least two reasons why this is
true. First, the fact that the renal disease is approaching end stage implies that
it has been a progressive disorder up until the time of diagnosis. Second,
processes of renal injury are thought to become self-perpetuating once
a suﬃcient amount of renal damage has occurred. In any event, animals with
advanced renal disease almost invariably experience further deterioration of
their renal function, although the rate of disease progression that occurs in
some individuals is more rapid than in others.

With the development of tests having the potential to detect mild renal

disease, however, a new problem arises, because one can no longer assume
that the detected nephropathy is (or will become) progressive. Many dogs and
cats that have some lesions in their kidneys never develop clinically
consequential renal dysfunction, much less renal failure. In dogs, for example,
several studies have found the prevalence of microscopic renal lesions in
randomly selected apparently healthy adult or aging dogs to be on the order of
40% to 80%, with most of the lesions observed being glomerular changes

. Renal lesions in apparently healthy dogs usually have limited

distribution (ie, they are focal rather than diﬀuse) and are mild or only
focally severe. It may be argued that the renal lesions in some of these dogs
would eventually progress to cause clinically evident renal disease or renal
failure. Nevertheless, this clearly does not happen in all dogs that have lesions
in their kidneys, because the proportion of all dogs that develop clinically
consequential renal disease during their lifetime is considerably less than 40%
to 80%.

As indicated previously, the whole point of early diagnosis of renal

disease is to permit institution of treatment to forestall progression of the
disease to more advanced stages. Thus, for animals with progressive renal
disease, detection of mild renal disease is potentially beneﬁcial. For animals
with mild nonprogressive renal disease, however, detection of their disease
could be detrimental if it prompts indiscriminate treatment. There is little or
nothing to be gained by treating mild nonprogressive renal disease; thus,
treatment of such animals would involve costs and possibly side eﬀects
without the justiﬁcation of potential beneﬁt. As a result, detection of mild
renal disease brings veterinarians to a new dilemma—how to tell animals
with progressive versus nonprogressive renal disease apart. Proven methods
for accomplishing this have not yet been established, but my suggestion is to
consider all animals with evidence of mild renal disease as being ‘‘at risk’’ for
progressive disease. These animals should be monitored more carefully than
animals that are not at such risk, and the crucial question is ‘‘what happens
next?’’ Animals that manifest worsening trends, even if the increments of
change are small, should be carefully investigated and treated aggressively.

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Animals that have mild but stable disease should be monitored at appro-
priate intervals.

Urine testing

Evaluation of a complete urinalysis, including examination of urine

sediment, is a key element in the clinical investigation of illness in all dogs
and cats that may have disease of or involving their urinary tracts. Similarly,
laboratory testing to screen for renal disease or failure in apparently healthy
animals also should start with a complete urinalysis for several important
reasons. Aside from its potential to aid in discovery of nonurinary disease (eg,
diabetes mellitus), a complete urinalysis is needed to screen for evidence of
subclinical excretory pathway disorders (eg, bacterial urinary tract infection,
urolithiasis) as well as to provide the necessary context for proper in-
terpretation of the results of tests (eg, indices of proteinuria and azotemia)
that are used to screen for early renal disease or failure. Subclinical excretory
pathway disorders not only occur with suﬃcient frequency to warrant eﬀorts
to detect and treat them in their own right but can incite renal disease and lead
to renal failure. Additionally, tests that detect excess amounts of protein in
the urine do not in themselves provide information about where in the urinary
tract or how the protein entered the urinary space. Lesions that generate
hemorrhage or exudation into the urine may cause plasma proteins, including
albumin, to be found in the urine regardless of whether the lesions are located
in the kidneys or the excretory pathway. The hallmark of such sources of
proteinuria is that the hemorrhage, inﬂammation, or both cause a coincident
ﬁnding of suﬃcient hematuria, pyuria, or both to account for the proteinuria.
Consequently, the ﬁrst step in interpreting any positive test result for
albuminuria or proteinuria needs to be evaluation of the results of a sediment
examination, preferably of the same urine specimen. In much the same way,
evaluation of the coincident urine speciﬁc gravity is a crucial initial step in
determining whether any increase in serum creatinine concentration that is
observed might have an entirely prerenal cause.

Proteinuria

Urine obtained from healthy dogs or cats with healthy kidneys typically

contains a small amount of protein, but as a diagnostic term, proteinuria
generally is taken to mean detection of an abnormal (ie, excessive) amount of
protein in the urine. Several diﬀerent methods to detect proteinuria can be
used to screen dogs and cats. These include semiquantitative tests performed
in a conventional urinalysis, determination of urine protein/creatinine ratio
(UPC), and assay of urine albumin concentration. Each of these methods has
its place in veterinary practice, none of the methods entirely replaces the
others, and they can be used in a complementary fashion.

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

A variety of conditions can cause the urine formed by healthy kidneys (ie,

kidneys that do not contain lesions) to contain excess protein. This type of
proteinuria has been classiﬁed as ‘‘functional renal proteinuria’’ (as opposed
to ‘‘pathologic renal proteinuria’’). The hallmark of such functional pro-
teinuria is that it is mild and transient, but pathologic proteinuria generally is
persistent unless the renal lesions that are responsible for it actually resolve.
To avoid undue concern about proteinuria that might be functional, it is
important to distinguish between transient proteinuria and persistent pro-
teinuria, especially when the magnitude of proteinuria is not large. The only
way to know whether proteinuria is transient or persistent, of course, is to
wait an appropriate interval and repeat the test. There is not a well-
established length of time to wait before retesting, and the most appropriate
interval might vary with circumstances; however, my recommendation is to
consider proteinuria that can be demonstrated repeatedly over a period of
1 month (eg, two tests 1 month apart) or more to be persistent. Moreover,
this principle applies regardless of what method(s) is(are) used to detect
the proteinuria. Persistent proteinuria should be considered a sign of renal
disease (once the possibility of it having a prerenal or postrenal origin has
been excluded properly), even if the magnitude of proteinuria (or albumin-
uria) is small.

Two types of semiquantitative tests for protein are common components

of a complete urinalysis; the reagent pad (dipstick) colorimetric test and the
sulfosalicylic acid (SSA) turbidimetric test. The amount of protein found in
the urine of normal cats and dogs lies below the lower limit of detection for
both of these tests; thus, normal animals should always test negative for
proteinuria by these methods. For technical reasons related to how dipstick
colorimetric tests work, however, they usually indicate a weak positive
reaction (trace to 1

þ) in moderately to well-concentrated canine and feline

urine specimens even when proteinuria is not present. Such erroneous
dipstick reactions occur because the buﬀer capacity of the reagent strip often
is insuﬃcient for the urine of dogs and cats, which frequently have urine that
is more concentrated than the urine of human beings for whom the test
strips were developed

. Misleading dipstick colorimetric tests for protein

can be distinguished from positive reactions that actually are indicative of
proteinuria by performing an SSA turbidimetric test on supernatant urine
from the same specimen

[14]

. The SSA turbidimetric test result will be

positive if the proteinuria is real and negative if it is not. Aside from these
spurious weak positive reactions in concentrated urine, dipstick colorimetric
test results are informative. Strong positive test results (eg, 2

þ, 3þ, 4þ)

usually are indicative of proteinuria, and negative test results are obtained
only when the protein content of the urine is below the detection limit for
the test. Many veterinary laboratories routinely perform an SSA turbidi-
metric test on every urine specimen, or at least on all that test positive for
protein by the dipstick colorimetric method (ie, to conﬁrm the result as cited
previously). Having been provided with results from both types of semi-

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

quantitative tests for most urine specimens that I have evaluated over nearly
three decades, my personal approach (and recommendation) is to focus
mainly on the SSA turbidimetric test result when interpreting the ﬁndings of
a routine urinalysis.

Consideration should also be given to urine speciﬁc gravity when

interpreting the results of semiquantitative tests for proteinuria. Any given
semiquantitative test result is likely to be associated with a larger amount of
proteinuria if the observed urine speciﬁc gravity is lower (ie, the urine is less
well concentrated) than if it is higher. Similarly, at any given urine speciﬁc
gravity value, a greater semiquantitative test result is likely to be associated
with a larger amount of proteinuria than if the test result was lower.
Regardless of these generalities, however, there is no reliable way to interpret
semiquantitative test results in conjunction with urine speciﬁc gravity values
to ascertain with conﬁdence the amount of protein being lost in the urine.
Therefore, the next step when a semiquantitative test, especially the SSA
turbidimetric test, gives a positive result is to determine the UPC value
regardless of the urine speciﬁc gravity.

The UPC value provides an index of total urinary protein loss. The

concentration of all urine proteins is determined and then divided by the
urine concentration of creatinine. Because the total amount of creatinine
excreted daily by an individual changes little, dividing the urine protein
concentration by the urine creatinine concentration eﬀectively adjusts the
protein concentration for variations in urine volume and concentration. This
permits UPC values to be used for comparison of total urinary protein loss at
diﬀerent times in a single animal as well as among animals. In veterinary
medicine, it is conventional to express both the urine protein concentration
and the urine creatinine concentration in the same units (ie, milligrams per
deciliter); thus, UPC values calculated using such concentrations are unitless
ratios. In human medicine, however, ratios of urine protein to creatinine are
sometimes expressed as milligrams of protein per gram of creatinine.

The UPC cutoﬀ that has been most widely used in dogs and cats to

diﬀerentiate values thought to be indicative of proteinuria from those that are
not is 1.0 (ie, with UPC values [1.0 being considered abnormal)

[15–18]

Numerous reports of UPC values determined in healthy young adult dogs
and cats indicate that such animals typically have a UPC value less than 0.5,
however

[16–23]

. Indeed, some authorities recommend interpreting UPC

values less than 0.5 as normal and those that are greater than or equal to 0.5
but less than or equal to 1.0 as questionable, with values greater than 1.0
considered abnormal

[13,24]

. Interpretation of UPC values is a prototypical

example of the need to select an appropriate cutoﬀ value for the way the test
result is being used. In dogs, for example, the cutoﬀ must be set as high as 2.0
to be conﬁdent that no dog is incorrectly labeled as proteinuric

[16,20]

especially if puppies less than 4 months of age are being evaluated

[25,26]

Dogs with a UPC value greater than or equal to 2.0 are certain to be abnormal
(ie, the cutoﬀ value yielding near-perfect speciﬁcity). Regardless of age,

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

however, healthy dogs do not have UPC values that are persistently greater
than or equal to 0.5. Therefore, a UPC value greater than or equal to 0.5 is the
more appropriate cutoﬀ to use when screening for proteinuria in apparently
healthy dogs, and the important issue that is ‘‘questionable’’ about UPC
values between 0.5 and 1.0 (or 2.0) is whether or not the proteinuria proves to
be persistent. Similarly, the most appropriate UPC cutoﬀ to use for cats
should not be higher than 0.5. Emerging evidence from recent studies of cats
with CKD suggests that even relatively small UPC increases (ie, into the 0.5–
1.0 range) are consequential because they are associated with greater risk of
more rapid disease progression

For animals that have a positive semiquantitative test for protein in

a urinalysis that does not reﬂect concomitant urinary hemorrhage or
inﬂammation, evaluation of the UPC value is the most logical next step.
Usually the UPC value conﬁrms the existence of proteinuria and provides
an index of the magnitude of proteinuria for comparison with subsequent
values as monitoring continues. If the positive semiquantitative test result
is not an indication of actual proteinuria (eg, as might be the case with
a weak positive reaction in a highly concentrated urine sample), the UPC
value is less than 0.5. For animals that unequivocally have renal pro-
teinuria (eg, dogs with a UPC value [2.0 that is otherwise unexplained),
testing to detect albuminuria is not worthwhile. The urine is sure to
contain an abundance of albumin, and the test result does not provide
additional diagnostically useful information. For animals with UPC values
in the ‘‘questionable’’ range, however, testing for albuminuria may be
helpful. Additionally, testing for albuminuria is the logical next step to
screen seemingly healthy animals (ie, those with negative semiquantitative
tests or UPC values \0.5) for evidence of renal disease. In my judgment,
however, it is not justiﬁable to screen all seemingly healthy animals for
albuminuria routinely. Seemingly healthy animals that should be screened
for albuminuria are those with an increased risk of having renal disease
either because of their age or some other factor (eg, a breed-associated
predisposition).

Assays have been developed to measure the concentration of albumin in

the urine of dogs and cats using species-speciﬁc antibody-mediated methods

[28,29]

. These assays have been calibrated to work optimally for measuring

low but nonetheless abnormal concentrations of albumin that usually would
escape detection by conventional semiquantitative tests (eg, dipstick color-
imetric tests). As with the concentration of all urine proteins used in the
UPC, the concentration of urine albumin must be adjusted in some way for
diﬀerences in urine volume and concentration so as to permit comparison of
values within and among individuals. Although it is possible to calculate
a urine albumin/creatinine ratio (milligrams of albumin/grams of creatinine)
for this purpose, the manufacturer of the commercially available tests has
chosen to adjust (ie, ‘‘normalize’’) concentrations of urine albumin to
a speciﬁc gravity of 1.010. When normalized to a urine speciﬁc gravity of

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

1.010, the concentration of albumin in the urine (nUAlb) of normal dogs and
cats is less than 1 mg/dL.

The term microalbuminuria refers to a particular kind and degree of

proteinuria, that is, an abnormal amount of albumin in a urine specimen that
would likely test negative for protein if evaluated by other more conventional
testing methods. Because the lower limit of urine protein detection by dip-
stick colorimetric tests is approximately 30 mg/dL, the nUAlb of dogs and
cats that is deﬁned as microalbuminuria is nUAlb greater than or equal to 1.0
mg/dL and less than 30 mg/dL. Greater concentrations of urine albumin (ie,
nUAlb [30 mg/dL) also are abnormal, but use of the micro preﬁx no longer
applies; that is, nUAlb greater than or equal to 30 mg/dL deﬁnes albuminuria
in much the same way that a UPC value greater than or equal to 0.5 deﬁnes
proteinuria. As discussed previously, however, it is not proteinuria (or
albuminuria), per se, that is a reliable indicator of renal disease; it is
persistent proteinuria or albuminuria (including microalbuminuria) other-
wise unexplained by prerenal or postrenal abnormalities that carries this
implication. Thus, screening for early renal disease by testing for proteinuria
requires repetitive urine tests to demonstrate that the proteinuria is persistent
as well as adequate ancillary testing to exclude nonrenal sources of the excess
proteins found in the urine.

Microalbuminuria, albuminuria, or proteinuria that is of renal origin

usually indicates the existence of altered permselectivity of the glomerular
ﬁltration barrier, although abnormalities of renal tubular function also can
be at least partially responsible (ie, as a result of inability of the renal tubules
to reabsorb proteins that normally traverse the glomerular capillary wall).
Mechanisms that alter glomerular permselectivity in dogs and cats generally
are set into motion initially by nonrenal disease processes. Discovery of an
underlying infectious, inﬂammatory, metabolic, or neoplastic condition,
especially if the condition is potentially treatable, likely is the single most
beneﬁcial outcome that may arise from detection of altered glomerular
permselectivity. For this reason, detection of microalbuminuria, albumin-
uria, or proteinuria should prompt an assiduous search for such an
underlying condition. Management dilemmas arise, however, when the
search for an underlying condition proves unproductive, especially when the
magnitude of microalbuminuria, proteinuria, or albuminuria is small and
renal function based on other indices (eg, urine speciﬁc gravity, serum
creatinine concentration) remains good. Because it is likely that at least
some and possibly many animals in this category have mild nonprogressive
renal disease for which treatment is not necessary, it seems to me most
prudent to carefully monitor such patients unless or until they manifest
some indication of a worsening trend (eg, an increasing magnitude of
proteinuria, diminishing urine-concentrating ability, increasing serum cre-
atinine concentration). In my judgment, the key is to be suﬃciently vigilant
to detect such indications of progressing renal injury in a timely manner if
they occur.

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Urine speciﬁc gravity

Observation of a suﬃciently high urine speciﬁc gravity value, greater than

1.035 in dogs and 1.040 in cats, always is evidence of ‘‘adequate’’ urine-
concentrating ability (ie, ability of the kidneys to concentrate the urine
suﬃciently well to indicate that renal function is adequate to maintain normal
homeostasis). The implications of ﬁnding urine speciﬁc gravity values less
than these cutoﬀs, however, depend on other concomitant ﬁndings. When
found together with either azotemia or a physiologic state that normally
causes healthy kidneys to form well-concentrated urine (eg, dehydration),
urine speciﬁc gravity values that are less than these adequate cutoﬀs (ie, are
less than should be expected) are an indication of impaired urine-concen-
trating ability. Moreover, the greater the disparity between the expected
value and the observed value, the more compelling is the evidence of renal
dysfunction. In the absence of evidence of a context in which formation of
well-concentrated urine should be expected, however, urine speciﬁc gravity
values below these cutoﬀs provide little or no information about urine-
concentrating ability. Depending on their state of water balance, healthy
animals with normal kidneys can excrete urine that is not well concentrated
or urine that is diluted compared with plasma. For this reason, it is not
possible to screen apparently healthy animals eﬃciently for renal dysfunction
using urine speciﬁc gravity values alone.

Nonetheless, consideration of the coincident urine speciﬁc gravity always

is an important part of interpreting the results of screening tests that are
useful, such as for proteinuria or azotemia. Therefore, the urine speciﬁc
gravity should be determined and recorded whenever a test to screen for renal
disease is performed.

Blood testing

Plasma creatinine concentration

Plasma creatinine concentration is determined by the balance between

daily endogenous creatinine production, which is proportional to muscle
mass, and daily excretion of creatinine. In dogs and cats, creatinine excretion
is accomplished almost exclusively by glomerular ﬁltration. Under steady-
state conditions, the total amount of creatinine made daily and the amount
excreted daily are equal whether or not renal function is normal. Because
day-to-day variation in muscle mass is relatively small, daily creatinine
production is fairly constant, especially in healthy dogs and cats. Changes in
muscle mass, however, such as loss of muscle mass associated with aging or
chronic illness, do alter daily creatinine production in individual animals over
the course of their life. Nonetheless, because daily creatinine production
usually is fairly constant, the main factor that causes plasma creatinine
concentration to change is GFR. Thus, changes in GFR reliably produce

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predictable changes in plasma creatinine concentration, but for the purposes
of early diagnosis and monitoring of renal disease, interpretation of plasma
creatinine concentration requires recognition of two important considera-
tions. The ﬁrst of these is that reference ranges for plasma creatinine
concentration in normal dogs and cats are relatively wide, which mostly
relates to the issue of screening apparently healthy animals for early renal
disease. The second is that meaningful changes in plasma creatinine
concentration are small in their absolute (but not relative) size during the
early stages of CKD, which mostly relates to the issue of monitoring animals
that are judged to be at risk for progressive renal disease.

In human beings, plasma creatinine concentration can be used to calculate

a suﬃciently accurate estimate of GFR for clinical patient care by taking the
factors of gender, age, race, and, sometimes, body size into account because
of their eﬀects on relative muscle mass

. Similarly, normal reference

ranges for plasma creatinine concentration in human beings diﬀer from one
another depending on such factors as gender and age; however, the ranges of
expected values are notably smaller than the commonly used ranges for such
values in dogs and cats. For examples, normal reference ranges given for
adult men (0.71–1.27 mg/dL) and women (0.60–1.15 mg/dL)

are not as

wide as those for dogs (0.5–1.4 mg/dL) and cats (0.7–2.2 mg/dL)

. The

existence of such wide reference ranges for animals is at least partly related to
the fact that individual animals with diverse, and not necessarily comparable,
relative muscle mass (as inﬂuenced by such factors as breed, size, gender, and
age) were included in the groups of normal animals used to establish the
range. As an example of the inﬂuence of body type on normal plasma
creatinine concentration, it has been observed that plasma creatinine
concentrations are higher in Greyhounds than in non-Greyhound breeds

. Another consideration is the possibility that some of the ‘‘apparently

healthy’’ animals used to establish the normal reference ranges might have
had unrecognized renal disease.

Because of the broad reference ranges used for dogs and cats, individual

animals can have a substantial decline in GFR and proportional increase in
plasma creatinine concentration without the value necessarily exceeding the
cutoﬀ that deﬁnes the upper limit of normal. For example, if a dog with
a normal plasma creatinine concentration of 0.9 mg/dL suﬀered a 35%
reduction of its GFR, the plasma creatinine concentration would only be
expected to increase to 1.4 mg/dL. This means that the ranges of plasma
creatinine concentrations in normal animals and in animals with their GFR
reduced by early renal disease overlap considerably. This impedes use of
plasma creatinine concentration to screen for early renal disease in dogs and
cats; however, it does not preclude eﬀective use of plasma creatinine
concentration to monitor individual animals for changes in their renal
function. Serial evaluations of plasma creatinine concentration in dogs with
declining GFR during early stages of progressive renal disease demonstrate
a recognizably abnormal trend of progressively increasing plasma creatinine

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

concentrations, albeit ‘‘within the normal range’’ and by small increments
initially (

The usefulness of applying plasma creatinine concentration as a screening

test for early renal disease in apparently healthy animals might possibly be
improved by lowering the cutoﬀ value. In dogs, for example, lowering the
cutoﬀ used to deﬁne ‘‘abnormal’’ from greater than or equal to 1.4 mg/dL to
greater than or equal to 1.3 mg/dL would improve sensitivity (ie, ﬁnd more
true-positive dogs with renal disease) but would also lower speciﬁcity (ie,
raise unwarranted concern about false-positive dogs that do not have renal
disease). It is rational (but unproven insofar as I know) to expect that the
speciﬁcity of diagnostic conclusions about animals with borderline or
slightly increased plasma creatinine concentrations (eg, in the range of
1.3–1.5 mg/dL using the example given previously) would be improved by
interpreting them in light of concurrent evaluations for proteinuria (in-
cluding microalbuminuria) and urine-concentrating ability. A dog that has
a plasma creatinine concentration of 1.4 mg/dL and also has persistent
proteinuria or repeated urine speciﬁc gravity values that are inadequately
concentrated is more likely to be an individual with early CKD than to be
healthy. My personal approach is not merely to expect healthy animals to
have plasma creatinine concentrations ‘‘within the reference range,’’ but to

2.0

4.0

2.0

4.0

6.0

Months of Age

Plasma Creatinine

Concentration (mg/dL)

GFR

P Creat

Glomerular Filtration Rate (ml/min/kg)

Fig. 2. Plots of plasma creatinine concentrations determined weekly and glomerular ﬁltration
rate (GFR) determined monthly by renal scintigraphy in a dog with a juvenile-onset progressive
renal disease (ie, X-linked hereditary nephropathy). Note that a trend of rising plasma
creatinine concentrations, albeit initially with small increments and through the upper portion
of the normal reference range, becomes apparent as soon as the dog’s GFR begins decreasing.
Moreover, GFR values below the reference range (ie, \2.5 mL/kg/min) do not prevail before
borderline to mild azotemia (plasma creatinine concentration: 1.6–2.0 mg/dL) develops. For
monitoring progression of the dog’s renal disease, serial plasma creatinine determinations were
as informative as serial GFR determinations.

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

expect most healthy animals to have values in the middle of that range. I
then take note of all values that are not ‘‘midrange normal’’ and ask myself
two questions. First, is there any concomitant ﬁnding (eg, proteinuria, low
urine speciﬁc gravity) that raises further concern? Second, is there any
concomitant ﬁnding (eg, muscular body type, highly concentrated urine that
might be associated with subclinical dehydration) that might explain the
high-normal creatinine value and thus dispel further concern?

Regardless of how animals are identiﬁed as being at risk for progressive

CKD (ie, screening for proteinuria, noting low urine speciﬁc gravity values,
observing ‘‘borderline or slightly increased’’ plasma creatinine concentra-
tions), the key question is ‘‘what happens next?’’ For monitoring animals at
risk for progressive CKD, serial evaluations of plasma creatinine are quite
useful. The animal is compared with itself rather than with other animals, and
even subtle trends can be identiﬁed. To be optimally useful, however, the
accuracy and repeatability of the method used to assay plasma creatinine
concentration must be high

[34,35]

. To minimize diﬃculties arising from

diﬀerences in calibration between laboratories, the best practice is to use
a single laboratory with good quality control to obtain the serial determi-
nations of plasma creatinine concentration. When variation attributable to
measurement inconsistencies is minimized, trends that are identiﬁed by serial
evaluations of plasma creatinine concentration during early stages of CKD
usually can be attributed to renal rather than extrarenal causes. This is
because the main extrarenal causes of variation of plasma creatinine
concentration in animals with reduced GFR, which are loss of muscle mass
and alterations of ﬂuid balance leading to additional prerenal impairments of
GFR, are minimal in their frequency of occurrence and in their relative
impact on plasma creatinine concentration in early versus later stages of
CKD. Therefore, although the magnitudes of change in plasma creatinine
concentration that occur during early stages of progressive CKD are small,
the changes that are observed are more highly likely to be caused by actual
changes in the animal’s intrinsic renal function than is the case later in the
course of the disease.

Clearance-based estimates of glomerular ﬁltration rate

The GFR generally is determined by measuring the renal clearance of an

appropriate compound. For any compound, renal clearance is the hypothetic
volume of plasma that must be completely cleared of its content of that
compound to account for the amount of the compound that is excreted by the
kidneys during a speciﬁc period. Key attributes of a compound that can be
used to estimate GFR are that it is not metabolized as it circulates in the
body, that it is freely ﬁltered through the glomerular ﬁltration barrier, and
that it is not reabsorbed or secreted by renal tubules. Inulin is the prototypical
material used to measure GFR, but other compounds that have been used to
make clearance-based estimates of GFR in dogs and cats include creatinine,

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G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

iohexol, and technetium Tc 99m–labeled diethylenetriaminepentaacetic acid
(DTPA)

[36–55]

. Depending on the method used, the amount of the

compound excreted by the kidneys is determined by assessing either the rate
of appearance of the compound in the urine or the rate of disappearance of
the compound from the plasma. Methods that use plasma disappearance
must account for disappearance of the compound from the plasma by
mechanisms other than by renal excretion (eg, diﬀusion into other body ﬂuid
compartments) and thus involve application of an appropriate pharmacoki-
netic model (one-compartment, two-compartment, or noncompartment
method) for analysis of the plasma disappearance curve

Every method for making clearance-based estimates of GFR is encum-

bered by various diﬃculties that limit its widespread use. First, there is the
issue of obtaining accurate assays of the compound being used. Except for
creatinine, assays for compounds used to measure GFR are cumbersome or
not widely available. Additionally, use of radiopharmaceutic agents requires
special expertise and regulatory approval. Methods that measure the urinary
appearance of a compound require timed urine collections that usually
involve urinary catheterizations, which may induce bacterial urinary tract
infections. Methods that measure plasma disappearance of a compound
require timed blood collections, and the accuracy of such methods generally
improves as more rather than fewer sample collection times are used to
construct the plasma disappearance curve. Moreover, every method esti-
mates GFR only during the time while the test is conducted and does not
discriminate whether or to what extent the GFR value is altered by prerenal
versus renal abnormalities. Interpretation of the resultant GFR value also
requires an appreciation of the repeatability of the test method

. Some

variation in results obtained within or between individual animals may be
caused by characteristics of the test rather than by diﬀerences in their actual
GFR values.

In my judgment, clearance-based estimates of GFR are most useful for the

investigation of individual animals in certain situations rather than for
widespread screening or monitoring of early renal disease and failure.
Examples of such situations include the investigation of polyuria in animals
with midrange normal plasma creatinine concentrations as well as the
investigation of animals with borderline increased plasma creatinine con-
centrations for which prior values are not available for comparison.

References

[1] Brown SA, Crowell WA, Barsanti JA, White JV, Finco DR. Beneﬁcial eﬀects of dietary

mineral restriction in dogs with marked reduction of functional renal mass. J Am Soc
Nephrol 1991;1(10):1169–79.

[2] Brown SA, Brown CA, Crowell WA, Barsanti JA, Allen T, Cowell C, et al. Beneﬁcial

eﬀects of chronic administration of dietary omega-3 polyunsaturated fatty acids in dogs
with renal insuﬃciency. J Lab Clin Med 1998;131(5):447–55.

882

G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

[3] Finco DR, Brown SA, Crowell WA, Groves CA, Duncan JR, Barsanti JA. Eﬀects of

phosphorus/calcium-restricted and phosphorus/calcium-replete 32% protein diets in dogs
with chronic renal failure. Am J Vet Res 1992;53(1):157–63.

[4] Grauer GF, Greco DS, Getzy DM, Cowgill LD, Vaden SL, Chew DJ, et al. Eﬀects of

enalapril versus placebo as a treatment for canine idiopathic glomerulonephritis. J Vet
Intern Med 2000;14(5):526–33.

[5] Jacob F, Polzin DJ, Osborne CA, Allen TA, Kirk CA, Neaton JD, et al. Clinical

evaluation of dietary modiﬁcation for treatment of spontaneous chronic renal failure in
dogs. J Am Vet Med Assoc 2002;220(8):1163–70.

[6] Adams LG, Polzin DJ, Osborne CA, O’Brien TD, Hostetter TH. Inﬂuence of dietary

protein/calorie intake on renal morphology and function in cats with 5/6 nephrectomy.
Lab Invest 1994;70(3):347–57.

[7] Barber PJ, Rawlings JM, Markwell PJ, Elliott J. Eﬀect of dietary phosphate restriction on

renal secondary hyperparathyroidism in the cat. J Small Anim Pract 1999;40(2):62–70.

[8] Brown SA, Brown CA, Jacobs G, Stiles J, Hendi RS, Wilson S. Eﬀects of the angiotensin

converting enzyme inhibitor benazepril in cats with induced renal insuﬃciency. Am J Vet
Res 2001;62(3):375–83.

[9] Brown SA, Finco DR, Brown CA, Crowell WA, Alva R, Ericsson GE, et al. Evaluation of

the eﬀects of inhibition of angiotensin converting enzyme with enalapril in dogs with
induced chronic renal insuﬃciency. Am J Vet Res 2003;64(3):321–7.

[10] Finco DR, Brown SA, Brown CA, Crowell WA, Cooper TA, Barsanti JA. Progression of

chronic renal disease in the dog. J Vet Intern Med 1999;13(6):516–28.

[11] Rouse BT, Lewis RJ. Canine glomerulonephritis: prevalence in dogs submitted at random

for euthanasia. Can J Comp Med 1975;39(4):365–70.

[12] Short RP, Lobetti RG, Nesbit JW. Renal pathology in working dogs in the South African

National Defence Force. J S Afr Vet Assoc 1999;70(4):158–60.

[13] Osborne CA, Stevens JB. Urinalysis: a clinical guide to compassionate patient care.

Shawnee Mission, KS: Bayer Corp., Agricultural Division, Animal Health; 1999.

[14] Moore FM, Brum SL, Brown L. Urine protein determination in dogs and cats: comparison

of dipstick and sulfasalicylic acid procedures. Vet Clin Pathol 1991;20(4):95–7.

[15] Fettman MJ. Comparison of urinary protein concentration and protein/creatinine ratio vs

routine microscopy in urinalysis of dogs: 500 cases (1987–1988). J Am Vet Med Assoc
1989;195(7):972–6.

[16] McCaw DL, Knapp DW, Hewett JE. Eﬀect of collection time and exercise restriction on

the prediction of urine protein excretion, using urine protein/creatinine ratio in dogs. Am J
Vet Res 1985;46(8):1665–9.

[17] Grauer GF, Thomas CB, Eicker SW. Estimation of quantitative proteinuria in the dog,

using the urine protein-to-creatinine ratio from a random, voided sample. Am J Vet Res
1985;46(10):2116–9.

[18] White JV, Olivier NB, Reimann K, Johnson C. Use of protein-to-creatinine ratio in a single

urine specimen for quantitative estimation of canine proteinuria. J Am Vet Med Assoc
1984;185(8):882–5.

[19] Finco DR, Brown SA, Brown CA, Crowell WA, Sunvold G, Cooper TL. Protein and

calorie eﬀects on progression of induced chronic renal failure in cats. Am J Vet Res 1998;
59(5):575–82.

[20] Center SA, Wilkinson E, Smith CA, Erb H, Lewis RM. 24-Hour urine protein/

creatinine ratio in dogs with protein-losing nephropathies. J Am Vet Med Assoc 1985;
187(8):820–4.

[21] Horauf A, Reusch C, Minkus G. [The diagnostic signiﬁcance of the protein-creatinine ratio

in urine for the diﬀerentiation of feline nephropathies]. Tierarztl Prax 1990;18(4):423–5.

[22] Monroe WE, Davenport DJ, Saunders GK. Twenty-four hour urinary protein loss in

healthy cats and the urinary protein-creatinine ratio as an estimate. Am J Vet Res 1989;
50(11):1906–9.

883

G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

[23] Adams LG, Polzin DJ, Osborne CA, O’Brien TD. Correlation of urine protein/creatinine

ratio and twenty-four-hour urinary protein excretion in normal cats and cats with
surgically induced chronic renal failure. J Vet Intern Med 1992;6(1):36–40.

[24] Lulich JP, Osborne CA. Interpretation of urine protein-creatinine ratios in dogs with

glomerular and nonglomerular disorders. Comp Vet Educ Pract Vet 1990;12:59–72.

[25] Lees GE, Helman RG, Kashtan CE, Michael AF, Homco LD, Millichamp NJ, et al. A

model of autosomal recessive Alport syndrome in English cocker spaniel dogs. Kidney Int
1998;54(3):706–19.

[26] Lees GE, Helman RG, Kashtan CE, Michael AF, Homco LD, Millichamp NJ, et al. New

form of X-linked dominant hereditary nephritis in dogs. Am J Vet Res 1999;60:373–83.

[27] Syme HM, Elliott J. Relation of survival time and urinary protein excretion in cats with

renal failure and/or hypertension [abstract]. J Vet Intern Med 2003;17:405.

[28] Syme HM, Elliott J. Development and validation of an enzyme linked immunosorbent

assay for the measurement of albumin in feline urine [abstract]. J Vet Intern Med 2000;14:
352.

[29] Pressler BM, Vaden SL, Jensen WA, Simpson D. Detection of canine microalbuminuria

using semiquantitative test strips designed for use with human urine. Vet Clin Pathol 2002;
31(2):56–60.

[30] Levey AS, Coresh J, Balk E, Kausz AT, Levin A, Steﬀes MW, et al. National Kidney

Foundation practice guidelines for chronic kidney disease: evaluation, classiﬁcation, and
stratiﬁcation. Ann Intern Med 2003;139(2):137–47.

[31] Couchoud C, Pozet N, Labeeuw M, Pouteil-Noble C. Screening early renal failure: cut-oﬀ

values for serum creatinine as an indicator of renal impairment. Kidney Int 1999;55(5):
1878–84.

[32] Tvedten H. Appendix II; reference values. In: Willard MD, Tvedten H, editors. Small

animal clinical diagnosis by laboratory methods. St. Louis: Saunders; 2004. p. 417–9.

[33] Feeman WE III, Couto CG, Gray TL. Serum creatinine concentrations in retired racing

Greyhounds. Vet Clin Pathol 2003;32(1):40–2.

[34] Coresh J, Astor BC, McQuillan G, Kusek J, Greene T, Van Lente F, et al. Calibration and

random variation of the serum creatinine assay as critical elements of using equations to
estimate glomerular ﬁltration rate. Am J Kidney Dis 2002;39(5):920–9.

[35] Coresh J, Eknoyan G, Levey AS. Estimating the prevalence of low glomerular ﬁltration

rate requires attention to the creatinine assay calibration. J Am Soc Nephrol 2002;13(11):
2811–2.

[36] Krawiec DR, Badertscher RR, Twardock AR, Rubin SI, Gelberg HB. Evaluation of

99mTc-diethylenetriaminepentaacetic acid nuclear imaging for quantitative determination
of the glomerular ﬁltration rate of dogs. Am J Vet Res 1986;47(10):2175–9.

[37] Gleadhill A, Michell AR. Evaluation of iohexol as a marker for the clinical measurement

of glomerular ﬁltration rate in dogs. Res Vet Sci 1996;60(2):117–21.

[38] Kampa N, Bostrom I, Lord P, Wennstrom U, Ohagen P, Maripuu E. Day-to-day

variability in glomerular ﬁltration rate in normal dogs by scintigraphic technique. J Vet
Med A Physiol Pathol Clin Med 2003;50(1):37–41.

[39] Heiene R, Moe L. Pharmacokinetic aspects of measurement of glomerular ﬁltration rate in

the dog: a review. J Vet Intern Med 1998;12(6):401–14.

[40] Haller M, Muller W, Binder H, Estelberger W, Arnold P. Single-injection inulin

clearance—a simple method for measuring glomerular ﬁltration rate in dogs. Res Vet
Sci 1998;64(2):151–6.

[41] Bovee KC, Joyce T. Clinical evaluation of glomerular function: 24-hour creatinine

clearance in dogs. J Am Vet Med Assoc 1979;174(5):488–91.

[42] Finco DR, Coulter DB, Barsanti JA. Simple, accurate method for clinical estimation of

glomerular ﬁltration rate in the dog. Am J Vet Res 1981;42(11):1874–7.

[43] Labato MA, Ross LA. Plasma disappearance of creatinine as a renal function test in the

dog. Res Vet Sci 1991;50(3):253–8.

884

G.E. Lees / Vet Clin Small Anim 34 (2004) 867–885

[44] Watson ADJ, Lefebvre HP, Concordet D, Laroute V, Ferre JP, Braun JP, et al. Plasma

exogenous creatinine clearance test in dogs: comparison with other methods and proposed
limited sampling strategy. J Vet Intern Med 2002;16(1):22–33.

[45] Brown SA, Haberman C, Finco DR. Use of plasma clearance of inulin for estimating

glomerular ﬁltration rate in cats. Am J Vet Res 1996;57(12):1702–5.

[46] Brown SA, Finco DR, Boudinot FD, Wright J, Taver SL, Cooper T. Evaluation of a single

injection method, using iohexol, for estimating glomerular ﬁltration rate in cats and dogs.
Am J Vet Res 1996;57(1):105–10.

[47] Finco DR, Braselton WE, Cooper TA. Relationship between plasma iohexol clearance and

urinary exogenous creatinine clearance in dogs. J Vet Intern Med 2001;15(4):368–73.

[48] Moe L, Heiene R. Estimation of glomerular ﬁltration rate in dogs with 99M-Tc-DTPA and

iohexol. Res Vet Sci 1995;58(2):138–43.

[49] Barthez PY, Hornof WJ, Cowgill LD, Neal LA, Mickel P. Comparison between the

scintigraphic uptake and plasma clearance of 99mTc-diethylenetriaminepentaacetic acid
(DTPA) for the evaluation of the glomerular ﬁltration rate in dogs. Vet Radiol Ultrasound
1998;39(5):470–4.

[50] Gleadhill A, Peters AM, Michell AR. A simple method for measuring glomerular ﬁltration

rate in dogs. Res Vet Sci 1995;59(2):118–23.

[51] Barthez PY, Chew DJ, DiBartola SP. Simpliﬁed methods for estimation of 99mTc-

pentetate and 131I-orthoiodohippurate plasma clearance in dogs and cats. J Vet Intern
Med 2001;15(3):200–8.

[52] Kampa N, Wennstrom U, Lord P, Twardock R, Maripuu E, Eksell P, et al. Eﬀect of

region of interest selection and uptake measurement on glomerular ﬁltration rate measured
by 99mTc-DTPA scintigraphy in dogs. Vet Radiol Ultrasound 2002;43(4):383–91.

[53] Barthez PY, Chew DJ, DiBartola SP. Eﬀect of sample number and time on determination

of plasma clearance of technetium Tc 99m pentetate and orthoiodohippurate sodium I 131
in dogs and cats. Am J Vet Res 2000;61(3):280–5.

[54] Uribe D, Krawiec DR, Twardock AR, Gelberg HB. Quantitative renal scintigraphic

determination of the glomerular ﬁltration rate in cats with normal and abnormal kidney
function, using 99mTc-diethylenetriaminepentaacetic acid. Am J Vet Res 1992;53(7):
1101–7.

[55] Rogers KS, Komkov A, Brown SA, Lees GE, Hightower D, Russo EA. Comparison of

four methods of estimating glomerular ﬁltration rate in cats. Am J Vet Res 1991;52(6):
961–4.

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Renal biopsy: methods and interpretation

Shelly L. Vaden, DVM, PhD

Department of Clinical Sciences, North Carolina State University, College of Veterinary

Medicine, 4700 Hillsborough Street, Raleigh, NC 27606, USA

Renal diseases are common in dogs and cats. Historical information,

physical examination, and clinical laboratory data often allow for the
diﬀerentiation of renal diseases into the general categories of acute renal
failure, chronic renal failure, and glomerular disease. Renal biopsy is often
required to establish a deﬁnitive diagnosis and determine the severity of the
lesion. A precise and accurate histologic diagnosis may also be needed to
formulate an optimal treatment plan. Accurate assessment of response to
therapy requires knowledge of the type and severity of the disease being
treated

[1–4]

There is often reluctance on the part of the practitioner to pursue renal

biopsy in the clinical evaluation of patients. Many concerns probably
contribute to this reluctance, including the potential complications of renal
biopsy, the expenses associated with procurement of the renal biopsy
specimen and adequate evaluation of the renal biopsy specimen, and the
belief that the rendered diagnoses may lack consistency. Studies have shown
that the frequency of severe complications from renal biopsy is relatively
low and that renal biopsy minimally aﬀects renal function when proper
technique is employed

[1,2,5–11]

. The expense of the renal biopsy procedure

can be minimized by correct patient selection and the use of proper
technique. Consistent and accurate diagnoses are more likely to be obtained
when renal biopsy specimens are appropriately processed and evaluated.
The purpose of this article is to discuss patient selection and evaluation,
renal biopsy techniques, expected complications of renal biopsy, and
appropriate processing and evaluation of the renal biopsy specimen.

Patient selection

Renal biopsy is indicated only when the results are likely to alter patient

management by providing an accurate histologic diagnosis or by facilitating
prognostication. Patients whose management is most likely to be altered by

E-mail address:

shelly_vaden@ncsu.edu

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.010

Vet Clin Small Anim

34 (2004) 887–908

results of a renal biopsy include those with glomerular disease (protein-
losing nephropathy) or acute renal failure. Client factors that also need to be
considered include the ability to incur the expense of the procedure and
proper evaluation of the specimen as well as the desire to pursue additional
treatment of a dog or cat as may be indicated once an accurate histologic
diagnosis is rendered.

Renal biopsy provides a deﬁnitive diagnosis of glomerular disease but

may not be needed if a potential underlying disease is identiﬁed and
treatment of this disease leads to resolution of proteinuria. Likewise,
a deﬁnitive diagnosis is unlikely to be of beneﬁt to the patient with
glomerular disease if end-stage renal disease is already present. In patients
that do not have end-stage renal disease, clinical decisions regarding
diagnosis, therapy, and prognosis can be made from the information
obtained through renal biopsy. In fact, obtaining an accurate histologic
diagnosis may be one of the more important factors in successful manage-
ment of the dog or cat with glomerular disease. In most cases, appropriate
evaluation of a renal biopsy specimen from a dog or cat with glomerular
disease includes light, electron, and immunoﬂuorescent microscopy.

Renal biopsy is indicated in the dog or cat with acute renal failure that is

either persistently severe (ie, persistent oliguria or uremia) or has de-
teriorated despite appropriate medical management. In these patients,
determining the speciﬁc cause of renal damage may lead to additional
therapeutic measures that are speciﬁc for the primary disease. Renal biopsy
can also facilitate prognostication in patients with acute renal failure
through assessment of the overall appearance of the tissue as well as
determination of the integrity of the tubular basement membrane. Light
microscopy may be all that is required in the evaluation of a biopsy
specimen from a patient with acute renal failure, but renal tissue should also
be collected for electron and immunoﬂuorescent microscopy in the event
that a glomerular disease is causing the acute renal failure.

Renal biopsy is unlikely to alter the prognosis, therapy, or outcome of

a patient with chronic renal failure. For patients with end-stage renal
disease, it is unlikely that the cause of the renal disease will be determined by
renal biopsy. Furthermore, studies in people undergoing renal biopsy have
demonstrated an increased risk of complication in patients with chronic
renal failure

. For these reasons, renal biopsy of patients with chronic

renal failure is generally not indicated.

Considerations before renal biopsy

Before renal biopsy, it is generally recommended that the patient be

thoroughly evaluated by obtaining a current history; performing a complete
physical examination; measuring systemic blood pressure; and analyzing
results of a biochemical proﬁle, complete blood cell count, urinalysis, and

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

coagulation proﬁle. These are performed not only to assess the current state
of health and suitability of the patient to undergo renal biopsy but to verify
that there are no contraindications to biopsy. Contraindications to renal
biopsy include the presence of an uncorrectable coagulopathy, severe
anemia, hydronephrosis, uncontrolled hypertension, large or multiple renal
cysts, perirenal abscess, extensive pyelonephritis, and end-stage renal
disease. Some authors have included a solitary kidney as a contraindication
to biopsy; however, biopsy of a solitary kidney may be performed if proper
technique is used and other contraindications are not present.

To identify patients with bleeding tendencies, a coagulation proﬁle is

generally recommended before renal biopsy. Studies in people have
demonstrated that preoperative coagulation proﬁles are not only unneces-
sary in patients without an obvious tendency to bleed but that abnormal
results do not correlate with bleeding after percutaneous liver biopsy or
general surgery

[13,14]

. Contrary to these ﬁndings, a recent report

demonstrated that dogs and cats with moderate to severe thrombocytopenia
(platelet counts \80,000 per microliter), dogs with prolonged one-stage
prothrombin time (OSPT [1.5

normal), and cats with prolonged activated

partial thromboplastin time (APTT [1.5

normal) had a greater risk of

hemorrhage from ultrasound-guided biopsy procedures

. Other patient

factors that may be associated with an increased risk of hemorrhage include
severe azotemia (serum creatinine [5 mg/dL), uncontrolled systemic hyper-
tension, or administration of nonsteroidal anti-inﬂammatory drugs within the
previous 5 days

[12,16–18]

. These patient abnormalities may not be absolute

contraindications to renal biopsy, however. Rather, when renal biopsy is
planned for patients with identiﬁed bleeding tendencies, the clinician should
be prepared to monitor the patient for severe perirenal hemorrhage after
biopsy and have suitable blood products from a compatible donor available to
administer to the patient if needed.

Abdominal ultrasound is generally performed as part of the initial

evaluation in patients with acute renal failure or protein-losing nephropa-
thy. In addition to assessing the size, shape, contour, and internal
architecture of the kidney, abdominal ultrasound allows for the identiﬁca-
tion of many of the previously listed contraindications to renal biopsy.
Severe hydronephrosis is a contraindication to renal biopsy because of
concern about penetrating the distended renal pelvis, which is likely to be
under increased pressure, as well as the increased risk of transecting the
larger arteries located in the corticomedullary junction and medulla.
Rupture of renal cysts and release of their contents into perirenal tissues
is associated with pain in people. Although the risk of inducing infection
through renal biopsy is low, cyst infections may be diﬃcult to treat because
of poor antibiotic penetration into the cyst ﬂuid. Concern over inducing
renal pain or infection via biopsy of renal cysts and the limited diagnostic
yield of a biopsy specimen that contains large cysts has led to the
recommendation that renal biopsy not be performed in kidneys that contain

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

large or multiple cysts. Perirenal abscessation and extensive pyelonephritis
are contraindications to renal biopsy because of the possibility that the
abdomen could become seeded with bacteria or other infectious agents.
Ideally, urinary tract infections should be eliminated before renal biopsy.

Procurement of the renal biopsy specimen

Renal biopsies can be obtained percutaneously (via laparoscopy, the

keyhole technique, ultrasound guidance, or blindly) or surgically. Regardless
of the method selected, only cortical tissue should be obtained. Most
diagnoses can be made through evaluation of cortical tissue alone. The risk
of serious hemorrhage may increase as the kidney is penetrated more deeply,
because renal vessels progressively increase in size from the surface of the
kidney toward the renal pelvis. Biopsy of the renal medulla may also be
associated with an increased risk of creating large areas of infarction and
ﬁbrosis

[19,20]

. Whenever possible, the biopsy is taken from either the cranial

or caudal pole of the kidney, because it is easier to stay in cortical tissue over
a larger portion of the kidney within the poles. In the dog with generalized
renal disease, the right kidney is often preferred over the left kidney for renal
biopsy. The right kidney is more stable, because the caudate lobe of the liver
provides resistance to movement during the biopsy procedure. Conversely,
the left kidney is more movable during the biopsy procedure but can be
found in a more caudal position, providing easier access in some deep-
chested dogs. Feline kidneys are located more caudally in the abdomen
compared with canine kidneys. Both feline kidneys can be easily localized
and immobilized, making both equally suitable for renal biopsy.

Patient sedation or anesthesia

Proper biopsy technique requires patient immobilization. Failure to

immobilize the patient may increase the likelihood of the development of
serious complications after renal biopsy. In addition, providing general
anesthesia to the patient during the biopsy procedure has been associated
with procurement of a quality biopsy specimen

. Although general

anesthesia is most likely to produce complete immobilization of the patient,
some patients who are extremely ill may be immobilized by sedatives alone.
Incomplete anesthesia of the peritoneum is a disadvantage of using only
sedation and local anesthesia and can result in sudden abdominal movement
during the biopsy procedure

Needle selection

A variety of needles are available for percutaneous kidney biopsy. The

Tru-cut biopsy needle was once the needle of choice, largely because other
options were limited. These needles can be diﬃcult to use, however.
Improper technique can result in poor-quality biopsy specimens and

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

inadvertent trauma to the kidney. With the availability of spring-loaded
needles and biopsy guns, the Tru-cut needle cannot be routinely recom-
mended. Likewise, the Vim-Silverman needle is no longer routinely used for
renal biopsy.

We prefer disposable spring-loaded biopsy needles for renal biopsy (E-Z

Core Single Action Biopsy Device; Products Group International, Lyons,
CO) (

). These needles can easily be operated using only one hand and

are available in 14-, 16-, 18-, and 20-gauge sizes with lengths of 6, 9, or 15 cm.
An advantage of this needle lies in the throw mechanism. During biopsy, the
spring-loaded stylet is advanced ﬁrst and is visible by ultrasound. The cutting
cannula is activated when the operator fully depresses the plunger with
a thumb. The biopsy instrument does not move deeper into the tissue during
activation of the cannula. The biopsy needle can then be removed from the
animal and the specimen retrieved from the specimen notch.

Alternatively, some institutions use automatic spring-loaded biopsy guns

(eg, Bard Biopsy; C.R. Bard, Murray Hill, NJ) (

) that can be loaded

with disposable needles of appropriate gauges and lengths available from
a variety of manufacturers. We have found that the weight and size of the gun
can render the use of these more diﬃcult compared with the spring-loaded
biopsy needles. The operator needs to have large and strong hands to use the
gun easily single handedly. Theses guns may be less suitable for obtaining
kidney biopsies compared with biopsy of other organs because of the limited
control of the depth of biopsy. Activation of the needle causes the throw to
go beyond where the tip of the needle is placed at the beginning of the
procedure. This is a potential source of operator error and may be associated
with an increased risk of penetration of the needle into the medulla.

We use either a 16- or 18-gauge needle that is 9 cm in length for our kidney

biopsies. In one study, use of a 14-gauge biopsy needle was associated with
a greater likelihood that biopsy specimens contained medulla

Obtaining a good-quality biopsy specimen with limited damage to the

kidney requires the use of a sharp biopsy needle. Because needles become

Fig. 1. Disposable spring-loaded biopsy instrument that can be used for renal biopsy. (Courtesy
of

E-Z Core Single Action Biopsy Device; Products Group International, Lyons, CO; with

permission.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

dull after several biopsies are taken and the needles are relatively in-
expensive, reuse of these disposable needles is not generally recommended.

Percutaneous biopsy using ultrasound guidance

Percutaneous renal biopsy using ultrasound guidance has become the

renal biopsy method of choice for dogs that are larger than 5 kg and for all
cats that do not have other contraindications for renal biopsy

. With this

technique, ultrasound is used to identify and examine the kidneys and
subsequently to guide correct placement of the needle. The normal renal
cortex can easily be diﬀerentiated from the medulla because of the relative
hyperechogenicity of the cortex; diﬀerentiation may be more diﬃcult in
diseased kidneys. The patient is placed in left lateral recumbency for biopsy
of the right kidney or in right lateral recumbency for biopsy of the left
kidney. The hair over the biopsy site is removed, the skin is aseptically
prepared, and sterile coupling gel is applied. A sterile sleeve is placed over
the ultrasound probe. The kidneys are scanned for general examination of
the renal architecture and for selection of the biopsy site. Once the site of
entry for the biopsy needle is determined, a small stab incision is made
through the skin, allowing the needle to stay sharp as it easily passes
through the skin. The tip of the needle is guided to the renal capsule with
one hand while the probe is held with the other (

Figs. 3 and 4

). The tip of the

needle should be placed through the renal capsule before activation to
prevent sliding of the needle along the capsule and avoid tearing the capsule.
The biopsy is then taken using the method that is appropriate for the
selected needle, making sure that the needle remains in the renal cortex
(

). When using a percutaneous method to obtain a renal biopsy from

a patient with glomerular disease, at least two quality samples of renal
cortex should be obtained using either a 16- or 18-gauge needle; one cortical
biopsy may be all that is required from a patient with acute renal failure (see
section on evaluation of the renal biopsy specimen). Digital pressure should

Fig. 2. Automatic spring-loaded biopsy gun and needle that can be used for renal biopsy.
(Courtesey of Bard Biopsy; C.R. Bard, Murray Hill, NJ; with permission.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

be applied to the kidney transabdominally for approximately 5 minutes after
biopsy to minimize hemorrhage.

Although needle guides are available for the ultrasound probe, we do not

generally use these. The guides are probe speciﬁc and are not available for
all probes. The requirement of speciﬁc computer software for operation of
the guides makes their use rather expensive. In addition, some operators ﬁnd
that the guides are conﬁning.

Keyhole technique

The keyhole technique can be used in dogs if ultrasound guidance is not

available

. The dog is placed in left lateral recumbency for biopsy of

Fig. 3. Schematic demonstrating the correct and incorrect methods of directing the renal biopsy
needle. Note that the needle should remain in the renal cortex, preferably in either the cranial or
caudal pole. The needle should not cross the corticomedullary junction or enter either the renal
medulla or pelvis.

Fig. 4. Ultrasound-guided renal biopsy in a dog. Note that the probe is held in one hand while
the needle is held in the other hand.

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the right kidney. The dog’s back should be facing the surgeon. The hair in
the lumbar fossa is clipped, and the skin is aseptically prepared. An oblique,
paralumbar, 7.5- to 10-cm skin incision is made on a line that bisects the
angle between the last rib and the edge of the lumbar musculature (

). It

may be impossible to palpate the kidney if the incision is made too far
caudal or ventral. A large vascular muscle mass needs to be dissected if the
incision is made too far dorsal. Too cranial an incision may lead to puncture
of the intercostal artery. The muscle ﬁbers are separated along muscle
planes, and the peritoneum is incised. The peritoneal incision must be large
enough to allow for easy insertion of the surgeon’s index ﬁnger over the
caudal pole of the kidney. The index ﬁnger holds the kidney against the edge
of the lumbar musculature. The other hand inserts the biopsy needle
through a separate small stab incision in the lateral body wall. The biopsy
needle is guided into the peritoneal cavity, and the tip of the needle is
positioned at the surface of the kidney or stabbed just through the capsule,

Fig. 5. The ultrasonographic image of the dog in

. This image is used to guide the needle

to the kidney and through the cortex.

Fig. 6. Making the incision for insertion of a ﬁnger for percutaneous renal biopsy in the dog
using the keyhole technique. (Courtesy of J.A. Barsanti, DVM, MS, Athens, GA.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

making sure the angle is such that the needle passes only through renal
cortex. Care should be taken not to penetrate too deeply beyond the renal
capsule, because this reduces the amount of renal cortex in the biopsy
specimen. The needle-cutting mechanism is activated as appropriate for the
selected needle. Additional biopsy specimens can be obtained as required.
After the needle is withdrawn, digital pressure should be placed for at least
5 minutes where the needle penetrated the kidney to minimize hemorrhage.

Use of the keyhole technique has been associated with the creation of

artifact in the biopsy sample, possibly because of the frequent need to
displace the kidney a considerable distance before biopsy. Such artifact
includes congestion of the peritubular and glomerular capillaries and
extravasation of erythrocytes into the tubular lumina and Bowman’s space.
In one study, however, there were no diﬀerences in specimen quality
between samples obtained with the keyhole technique compared with those
obtained by laparoscopy

Blind or palpation technique

Performing renal biopsy with guidance by palpation is rarely done in

dogs because of the more cranial location of the kidneys and the fact that
canine kidneys can be diﬃcult to immobilize by palpation. Blind biopsy is
more frequently performed in feline kidneys, which are relatively more
caudal in position and can be more readily immobilized. Nonetheless, we
prefer ultrasound guidance for renal biopsies in cats, because ultrasound
aﬀords the ability to concurrently assess the renal architecture and establish
that contraindications to renal biopsy are not present (eg, polycystic kidney
disease) as well as the ability to guide the biopsy needle through the cortex.

Blind renal biopsy in cats is performed with the cat in lateral recumbency.

Either kidney is localized by palpation. The hair is clipped from the area
over the kidney, and the skin is aseptically prepared. A small stab incision is
made in the skin with a scalpel blade. The kidney is immobilized with one
hand. The other hand advances the needle through the stab incision and
directs it toward the cranial or caudal pole of the kidney (

Fig. 7

). The tip of

the needle is positioned at the surface of the kidney or stabbed just through
the capsule, making sure the angle is such that the needle passes only
through renal cortex. Penetrating too deeply beyond the renal capsule
reduces the amount of renal cortex in the biopsy specimen. The cutting
mechanism of the needle is then activated. Additional biopsy specimens can
be obtained if needed. Digital pressure should be applied to the kidney for
approximately 5 minutes after biopsy to minimize hemorrhage.

Laparoscopic biopsy

Laparoscopy is an endoscopic procedure that is performed under sterile

conditions and allows for visual examination of the peritoneal cavity after

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establishment of a pneumoperitoneum

[1,23]

. Laparoscopy oﬀers an

advantage over the percutaneous biopsy techniques in that it allows for
direct visualization and inspection of the kidneys through insertion of a rigid
endoscope and permits visual control of the biopsy. Visualization of the
kidneys before biopsy leads to a higher likelihood that diagnostic tissue will
be obtained during biopsy, particularly if focal lesions are present. Because
the abdominal incision is small, laparoscopy is less invasive and can be
performed more quickly than surgery, allowing for comparatively less
patient morbidity. Like surgery, other abdominal organs can be inspected
and biopsy specimens collected, if necessary, during laparoscopy, although
complete abdominal exploration is not possible. As with surgery, postbiopsy
hemorrhage can be monitored during laparoscopy; direct pressure can be
applied with the laparoscope or laparoscopic tools if needed. Laparoscopy
requires appropriate equipment and operator expertise. Contraindications
to laparoscopy include peritonitis, extensive abdominal adhesions, hernias,
obesity, coagulopathies, and operator inexperience

. Potential compli-

cations of laparoscopy include the creation of air emboli, pneumothorax, or
subcutaneous emphysema; the introduction of gas into a hollow viscus;
damage to internal organs with the Verres needle or trocar; and cardiac
arrest. Evacuation of the urinary bladder and colon before penetration into
the abdomen minimizes the chance that these organs are punctured.
Operator experience and attention to detail as well as the use of a surgically
placed port instead of the Verres needle may reduce the likelihood of
complications

. In a study of laparoscopic renal biopsy performed in 37

dogs and one cat, none of these complications were encountered.

Laparoscopy can be performed through a right lateral, left lateral, or

midline incision. The right kidney is readily visible in dogs in left lateral
recumbency (

. To separate the abdominal wall from the organs,

a pneumoperitoneum is created via injection of carbon dioxide through the
Verres needle or a surgically placed trocar and cannula unit. If the Verres

Fig. 7. Percutaneous kidney biopsy in the cat. The kidney is immobilized by one hand while the
other manipulates the biopsy needle. (Courtesy of J.A. Barsanti, DVM, MS, Athens, GA.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

needle technique is used, the trocar and cannula unit for the laparoscope are
then inserted through a small (1-cm) skin incision. Once the assembly is in the
abdomen, the trocar is removed from the cannula and replaced by the
laparoscope. The abdominal organs can be systematically inspected. A biopsy
needle can then be introduced into the abdomen through a separate site. The
renal biopsy is taken while observing the procedure through the laparoscope.

Surgical biopsy

Surgical biopsy may be the preferred method in dogs that are small (\5 kg)

or in animals that either have isolated areas in the kidney (eg, large cysts) that
need to be avoided during the biopsy procedure or are undergoing laparotomy
for another reason. Likewise, surgical biopsy may be safer in some animals
that have other listed contraindications to biopsy. A surgical wedge biopsy is
preferred over a surgical needle biopsy. One has more control over the depth of
biopsy and the volume of tissue collected when a wedge biopsy is obtained
compared with a needle biopsy. It is not surprising that surgical wedge biopsies
were ﬁve times more likely to be of good quality than surgical needle biopsies

. The surgical biopsy can be obtained through a paracostal incision if only

one kidney is to be examined and biopsied or through a cranial midline
abdominal incision if other intra-abdominal procedures are to be performed
or both kidneys need to be examined before biopsy

[21]

. The paracostal

incision is made with the patient in lateral recumbency

. The incision is

parallel and 2 cm caudal to the last rib. The oblique muscles are divided
between ﬁbers and retracted. The kidney is located after separating the
transverse abdominal muscle. The kidney can be elevated through the incision
by placing umbilical tape around both poles. In obese animals, exposure of the
kidney may be diﬃcult through the paracostal incision. After exposure

Fig. 8. Close-up laparoscopic view of the right kidney in a dog. The tip of a biopsy needle is in
the ﬁeld of view. Note that the biopsy needle is directed at a shallow angle to the capsule and
away from the renal pelvis. (From Grauer G. Laparoscopy of the urinary tract. In: Tams TR,
editor. Small animal endoscopy. St. Louis: Mosby; 1999. p. 430; with permission.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

through either type of incision, the kidney is immobilized with the thumb and
foreﬁnger before biopsy. A wedge-shaped incision is made through the capsule
and into the cortex (

Fig. 9

). Tissue forceps are used to gently lift the biopsy

wedge while the scalpel blade is used to sever any remaining attachments.
Monoﬁlament, absorbable suture material (4-0) in a simple continuous
pattern is used to close the renal capsule.

Postbiopsy care of the patient

Isotonic ﬂuids should be given liberally intravenously for several hours

after renal biopsy in amounts needed to produce diuresis. In theory, this
reduces the likelihood of obstructing clots forming in the renal pelvis or
ureter. To reduce the risk of serious hemorrhage, the patient should be kept

Fig. 9. Surgical kidney biopsy. (A) The kidney is immobilized between the thumb and
foreﬁnger. A wedge-shaped incision is made through the capsule and cortex. (B) Tissue forceps
are used to gently lift the biopsy sample. Any remaining attachments are severed with the
scalpel blade. (C) The capsule is closed with 4-0 monoﬁlament absorbable suture material.
Pressure is applied with the thumb and foreﬁnger to appose the edges during suturing.
(Courtesy of E.A. Stone, DVM, MS, MPP, Raleigh, NC.)

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relatively quiet and in the hospital for 24 hours after biopsy. The patient’s
packed cell volume should be evaluated 24 hours after biopsy or sooner if
a concern arises that major bleeding is occurring. Dogs should be walked
only on a leash for 72 hours after biopsy. The color of urine should be
monitored for several days after renal biopsy. Gross hematuria is common
after renal biopsy and usually resolves within 24 hours. Persistent gross
hematuria warrants re-evaluation of the kidneys and biopsy site.

Complications associated with renal biopsy

Results of clinical studies indicate that complications after renal biopsy

are limited but vary in frequency from 1% to 18%

[1,2,5–10,17]

(

). This

variation in complication rate is almost certainly a result of biopsy technique
as well as patient status at the time of biopsy. In one retrospective study,
factors associated with the development of complications in dogs included
patient age greater than 4 years, body weight less than 5 kg, and presence of
severe azotemia (serum creatinine [5 mg/dL)

. A hospital factor that

seemed to be associated with the development of complications was having
a radiologist or internist perform the biopsy. This association may have been
a result of the use of percutaneous methods to collect the renal biopsy or the
use of sedation or injectable anesthesia instead of general anesthesia.

Box 1. Reported complications of renal biopsy

Arteriovenous fistula formation
Biopsy of nonrenal tissue (eg, liver, adrenal gland, fat, muscle,

connective tissue, spleen)

Cyst formation
Death
Hemorrhage
Microscopic hematuria
Macroscopic hematuria
Perirenal hematoma
Intrarenal hematoma
Lacerated renal artery or vein
Intra-abdominal hemorrhage caused by laceration of other

organ or vessel

Hydronephrosis
Infarction and thrombosis
Infection
Scar formation and fibrosis

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Microscopic hematuria is an expected ﬁnding after renal biopsy, de-

veloping in approximately 20% to 70% dogs and cats

[2,17]

. Microscopic

hematuria is self-limiting, generally resolving within 48 to 72 hours of
biopsy. Macroscopic hematuria is less common, developing in approxi-
mately 1% to 4% of dogs and cats after renal biopsy. Small perirenal
hematomas are also common after renal biopsy if the kidney is examined
carefully by ultrasound. Severe hemorrhage, often severe enough to require
blood transfusion, was the most common reported complication in one
study, occurring in 9.9% of dogs and 16.9% of cats

. In addition to the

coagulation abnormalities previously discussed, uncontrolled hypertension,
uremia, the administration of nonsteroidal anti-inﬂammatory drugs, and the
use of improper technique may increase the risk of serious hemorrhage

[9,16,20]

. Hydronephrosis developing secondary to obstruction of the renal

pelvis or ureter by a blood clot is an uncommon complication of renal
biopsy. Death is also an uncommon complication, occurring in 3% or less of
dogs and cats undergoing renal biopsy

[9,17]

Histologic changes in renal tissue after biopsy have been well docu-

mented. Linear infarcts representing needle tracts associated with varying
amounts of atrophy and ﬁbrosis seem to be common after renal biopsy

Retention cysts can also be found in association with the needle tract and
probably form secondary to tubular obstruction

. Studies have demon-

strated correlations between severe renal parenchymal changes of hemor-
rhage, thrombosis, infarction, and ﬁbrosis and the presence of major renal
vessels or medulla in the biopsy specimen

[19,20]

. These ﬁndings emphasize

the need to direct the biopsy needle only through cortical tissue during the
biopsy procedure, avoiding the corticomedullary junction and medulla.

Despite histologic changes, renal biopsy seems to have minimal eﬀect on

renal function

[9,11]

. One could imagine that inﬂicting renal damage to an

animal with preexisting renal disease could contribute to progressive loss of
renal function, however. Obtaining multiple biopsies of the kidney does not
seem to produce more damage than biopsy with only a single pass provided
that the biopsy needle remains in cortical tissue

Processing the renal biopsy specimen

An adequate sample of cortex has a minimum of ﬁve glomeruli when

examined by light microscopy, although one glomerulus may be all that is
needed to make a deﬁnitive diagnosis in generalized glomerular diseases.
Microscopic examination of each specimen using 10-fold magniﬁcation
provides immediate veriﬁcation that adequate biopsy samples have been
obtained. Ideally, this is performed with a transilluminating microscope, but
a standard light microscope can also be used. When two samples are obtained
from a patient with glomerular disease, one sample should be placed in
formalin and the other should be divided into two smaller pieces containing

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glomeruli. One of the pieces is put into a ﬁxative suitable for electron
microscopy (eg, 4% formalin plus 1% glutaraldehyde in sodium phosphate
buﬀer), and the other piece is frozen for immunoﬂuorescent microscopy. An
alternative to freezing is to immerse the tissue in ammonium sulfate-N-
ethylmaleimide ﬁxative (ie, Michel’s solution), which preserves tissue-ﬁxed
immunoglobulins. Wedge biopsies should be divided in a similar fashion;
tissue for electron microscopy should be minced appropriately. Although
formalin ﬁxation may be all that is required for adequate histopathologic
evaluation of specimens from patients with acute renal failure, samples
should also be collected for electron and immunoﬂuorescent microscopy in
the event that a glomerular disease is causing the acute renal failure.

Thin sections (2–4 lm) of paraﬃn-embedded tissue should be used for

light microscopy, because standard sections of 5 to 6 lm are too thick for
adequate assessment of glomerular cellularity and capillary loop thickness

[27,28]

. Hematoxylin and eosin staining can be used for initial assessment of

the general appearance of the specimen. Periodic acid–Schiﬀ (PAS), which
stains glycoproteins, is the preferred stain of many nephropathologists and is
particularly useful in the demonstration of interstitial and glomerular
scarring and assessment of the glomerular basement membrane. Methena-
mine silver speciﬁcally stains the basement membrane of the tubules,
glomeruli, and Bowman’s capsule. Trichrome is useful for evaluation of the
mesangium and is also the best light microscopic stain for visualization of
immunoglobulins. Congo red can be used to demonstrate the presence of
amyloid. Although electron microscopy of renal biopsy specimens has not
been used frequently enough in dogs and cats to determine the diagnostic
merit in groups of patients with varying presentations, it seems to be most
helpful in the evaluation of people with renal hematuria, a familial history of
renal disease, or proteinuria with normal renal excretory function. A sample
should be collected for electron microscopy in nearly all dogs and cats
undergoing renal biopsy, because the exact diagnosis is most likely unknown.
Immunoﬂuorescent microscopy or immunohistochemistry should include
stains for IgM, IgG, IgA, and C3 at a minimum. Immunologic studies are
becoming more readily available at a variety of laboratories. The re-
quirement that samples from patients with glomerular disease be evaluated
by electron microscopy may necessitate that samples from such patients be
submitted to, and perhaps even collected at, a veterinary teaching hospital.

Evaluating the renal biopsy specimen

Whenever possible, specimens should be evaluated by a pathologist with

expertise in nephropathology. Although renal biopsy specimens from dogs
with acute renal failure may be adequately evaluated by light microscopy
alone, specimens from proteinuric dogs and cats should be evaluated by
light, electron, and immunoﬂuorescent microscopy. Limiting the evaluation

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

in these patients to light microscopy alone often allows for only a subjective
interpretation of the glomerular lesion with too much room for error.
Although there are many reports describing glomerular lesions in dogs and
cats, the use of diﬀerent nomenclature and diﬀerent morphologic criteria
among pathologists sometimes makes interpretations and comparisons of
the data diﬃcult at best. A standard classiﬁcation system for the
characterization of glomerular lesions in dogs and cats needs to be
embraced. The World Health Organization (WHO) criteria for the
classiﬁcation of human glomerulopathies have proven to be applicable to
glomerular diseases in dogs

. General acceptance of this system would

lead to a better understanding of the natural history, pathogenesis, and
response to treatment of the various glomerular diseases in dogs and cats.

Evaluation of the renal biopsy specimen by light microscopy

The normal kidney is composed of glomeruli, tubules, interstitium, and

vasculature. The normal glomerulus contains four to eight lobules; each
lobule is composed of capillaries supported on a centrilobular core of
mesangial matrix (

Fig. 10

[27,28]

. The glomerular basement membrane is

thin, delicate, PAS-positive, and argyrophilic (ie, capable of being impreg-
nated with silver). The glomerular capillary lumen is normally widely patent
and lined with eosinophilic endothelial cell cytoplasm. In any section, the
glomeruli can appear to be of any size because the glomeruli have been cut
in diﬀerent planes. There should only be one to two nuclei per mesangial cell
region. The tubules normally have a thick basement membrane and are

Fig. 10. Normal glomerulus from a dog. Note that the capillary lumens are widely patent and
the capillary loops are thin, often appearing discontinuous. Hypercellularity is not present.
(From Vaden SL. Glomerular disease. In: Ettinger SJ, Feldman EC, editors. Textbook of
veterinary internal medicine. 6th edition, Philadelphia: WB Saunders; 2004, in press; with
permission.)

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separated from each other by a ﬁne interstitial stroma that contains
capillaries. The normal interstitium is relatively inconspicuous.

The entire section should be evaluated under low power to exclude focal

space-occupying lesions (eg, granuloma, neoplasia) and to evaluate the
distribution of other lesions. When glomerular lesions are present and nearly
all glomeruli are aﬀected, the disorder is generalized. If less than half of the
glomeruli are aﬀected, the disorder is focal. When evaluating individual
glomeruli, the pattern is diﬀuse or global if the entire glomerulus is aﬀected.
The lesion is segmental or local if only a few lobules within aﬀected
glomeruli are involved.

In membranous glomerulopathy, the glomerular basement membrane

appears uniformly thickened and more rigid than normal (

Fig. 11

). Because

the subepithelial immune deposits do not become impregnated with silver,
spikes may be identiﬁed on the outside of the glomerular basement
membrane when a silver stain is used. Membranoproliferative glomerulo-
nephritis is diagnosed when there is a thickened glomerular basement
membrane and mesangial hypercellularity (more than three nuclei per
mesangial region) (

Fig. 12

). The glomerulus may become segmented or

lobular in appearance. Type I membranoproliferative glomerulonephritis,
also called mesangiocapillary glomerulonephritis, can have a ‘‘railroad’’
appearance to the glomerular basement membrane when evaluated by light
microscopy and is often induced by infectious diseases. Type II membra-
noproliferative glomerulonephritis is also called dense deposit disease and

Fig. 11. Glomerulus from a dog with membranous nephropathy. Note the thickened and rigid-
appearing capillary loops and the lack of hypercellularity. (Courtesy of J.L. Robertson, VMD,
PhD.)

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

can be diﬀerentiated from type I via electron microscopy. In people, the
dense deposits of type II membranoproliferative glomerulonephritis are not
believed to be immune deposits; type II membranoproliferative glomerulo-
nephritis is not associated with infectious diseases. Mesangioproliferative
glomerulonephritis may have several subcategories (eg, IgA nephropathy)
and is characterized by focal segmental to diﬀuse global mesangial cell
hyperplasia. Lupus nephritis can be associated with any glomerular
abnormality and, when active, often has an interstitial inﬁltrate of mixed
inﬂammatory cells and acute damage to the tubules. Crescents can loosely
be deﬁned as two or more layers of cells in Bowman’s space and are
suggestive of severe pathologic injury. Crescentic glomerulonephritis is
diagnosed when crescents are present in 50% or more of the glomeruli.
Focal segmental glomerulosclerosis is diagnosed when the patient has
proteinuria and glomerulosclerosis of only a few lobules within aﬀected
glomeruli and another glomerular lesion is not present to explain the
sclerosis (

Fig. 13

). Amyloidosis is easy to diagnose by expansion of the

mesangium and glomerular basement membrane by acellular eosinophilic
staining material that stains red by Congo red (

Fig. 14

). Amyloid deposits

are birefringent when viewed under polarized light.

Tubular atrophy appears as irregular tubular basement membrane

thickening and luminal collapse and may suggest long-standing disease or
a normal aging process. The tubules may be more widely separated than
normal, with ﬁbrous tissue and lymphocytes present between the tubules.
Hyaline droplets or protein reabsorption droplets can sometimes be seen in

Fig. 12. Glomerulus from a dog with membranoproliferative (mesangiocapillary) glomerulo-
nephritis. Note the thickened capillary loops and mesangial hypercellularity resulting in the
segmented and lobular appearance. (From Vaden SL. Glomerular disease. In: Ettinger SJ,
Feldman EC, editors. Textbook of veterinary internal medicine. 6th edition. Philadelphia: WB
Saunders; 2004, in press; with permission.)

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the renal tubular cells of patients with proteinuria. The tubules should be
examined for the presence of casts and foci of degeneration, regeneration,
and necrosis. The interstitium should be examined for edema, ﬁbrosis,
inﬂammation, or combinations of these. Expansion of the interstitium may
occur with tubular atrophy, edema, inﬂammatory inﬁltration, or ﬁbrosis.

Evaluation of the renal biopsy specimen by electron microscopy

Parietal epithelial cells, visceral epithelial cells, endothelial cells, and

mesangial cells comprise the normal glomerulus and can easily be identiﬁed

Fig. 14. Congo red–stained section showing typical birefringence of glomerular amyloid
deposits. (Courtesy of S.P. DiBartola, DVM.)

Fig. 13. A glomerulus from a dog with a lesion resembling focal segmental glomerulosclerosis.
Note the relatively normal appearance of the glomerular sections, which are not sclerotic. (From
Vaden SL. Glomerular disease. In: Ettinger SJ, Feldman EC, editors. Textbook of veterinary
internal medicine. 6th edition. Philadelphia: WB Saunders; 2004, in press; with permission.)

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by electron microscopy. The parietal epithelial cells are ﬂattened cells that
line the inner surface of Bowman’s capsule. The visceral epithelial cells, or
podocytes, line the outer surface of the capillary loops and rest on the
glomerular basement membrane. The podocytes are characterized by foot
processes and form the outermost layer of the capillary wall. The endothelial
cells line the inner surface of the capillary loops with the nuclei disposed
centrilobularly toward the mesangium. The fenestrations of the endothelial
cell cytoplasm can be visualized easily. The normal glomerular basement
membrane should be approximately the same thickness as the base of a foot
process turned 90(. The mesangial matrix contains an interwoven network
of microﬁlaments.

Minimal change disease is diagnosed when minimal changes are visualized

via light microscopy (ie, slight hypercellularity as evident by three to four
nuclei in the mesangial region) and marked foot process eﬀacement is noted
via electron microscopy (

Fig. 15

). Membranous glomerulopathy has four

ultrastructural stages that correlate with temporal evolution of the disease and
may predict therapeutic outcome; electron microscopy can be used to stage
patients with membranous nephropathy (

Fig. 16

). Electron microscopy can

also be used to identify the dense deposits in type II membranoproliferative
glomerulonephritis and to verify the presence of electron-dense, presumably
immune, deposits in other patients with glomerulonephritis. Whereas
veterinarians have classically been taught to look for these deposits within
the glomerular basement membrane, in dogs with glomerulonephritis, these
deposits are located more frequently within the mesangium. The glomerular
basement membrane varies in thickness and is split into a number of layers in
patients with hereditary nephritis

Fig. 15. Electron micrograph of a glomerular capillary loop in a dog with minimal change
disease. There is eﬀacement of the foot processes. (From Vaden SL. Glomerular disease. In:
Ettinger SJ, Feldman EC, editors. Textbook of veterinary internal medicine. 6th edition.
Philadelphia: WB Saunders; 2004, in press; with permission.)

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Evaluation of the renal biopsy specimen by immunoﬂuorescent microscopy

The normal kidney should not have positive immunoﬂuorescent or

immunohistochemical staining. Staining is positive in animals with immune-
mediated disease, however, where there is either in situ immune complex
formation, as is believed to occur with membranous glomerulopathy, or
deposition of circulating complexes, as may occur with type I membrano-
proliferative glomerulonephritis. Because true antiglomerular basement
membrane glomerulonephritis has not been documented in the dog, the
expected staining pattern is discontinuous, occurring either in the capillary
loops, giving a circular appearance, or in the mesangium.

Summary

Renal biopsy most often is indicated in the management of dogs and cats

with glomerular disease or acute renal failure. Renal biopsy can readily be
performed in dogs and cats via either percutaneous or surgical methods.
Care should be taken to ensure that proper technique is used. When proper
technique is employed and patient factors are properly addressed, renal
biopsy is a relatively safe procedure that minimally aﬀects renal function.
Patients should be monitored during the postbiopsy period for severe
hemorrhage, the most common complication. Accurate diagnosis of
glomerular disease, and therefore accurate treatment planning, requires
that the biopsy specimens not only be evaluated by light microscopy using
special stains but by electron and immunoﬂuorescent microscopy.

References

[1] Grauer GF, Twedt DC, Mero KN. Evaluation of laparoscopy for obtaining renal biopsy

specimens from dogs and cats. J Am Vet Med Assoc 1983;183:677–9.

[2] Minkus G, Reusch C, Ho¨rauf A, et al. Evaluation of renal biopsies in cats and

dogs—histopathology in comparison with clinical data. J Small Anim Pract 1994;35:465–72.

[3] Richards NT, Darby S, Howie AJ, et al. Knowledge of renal histology alters patient

management in over 40% of cases. Nephrol Dial Transplant 1994;9:1255–9.

[4] Vilafranca M, Wohlsein P, Trautwein G, et al. Histological and immunohistological

classiﬁcation of canine glomerular disease. Zentralbl Veterinarmed A 1994;41:599–610.

Fig. 16. Ultrastructural stages in the progression of membranous nephropathy. (Courtesy of
J.C. Jennette, DVM, MS.)

907

S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

[5] Jeraj K, Osborne CA, Stevens JB. Evaluation of renal biopsy in 197 dogs and cats. J Am

Vet Med Assoc 1982;181:367–9.

[6] Le´veille´ R, Partington BP, Biller DS, Miyabayshi T. Complications after ultrasound-

guided biopsy of abdominal structures in dogs and cats: 246 cases (1984–1991). J Am Vet
Med Assoc 1993;203:413–5.

[7] Hager DA, Nyland T, Fisher P. Ultrasound-guided biopsy of the canine liver, kidney and

prostate. Vet Radiol 1985;26:82–8.

[8] Groman RP, Bahr A, Berridge BR, Lees GE. Eﬀects of serial ultrasound-guided renal

biopsies on kidneys of healthy adolescent dogs. Vet Radiol Ultrasound 2004;451:62–9.

[9] Osborne CA. Clinical evaluation of needle biopsy of the kidney and its complications in the

dog and cat. J Am Vet Med Assoc 1971;158:1213–28.

[10] Wise LA, Allen TA, Cartwright M. Comparison of renal biopsy techniques in dogs. J Am

Vet Med Assoc 1989;195:935–9.

[11] Drost WT, Henry GA, Meinkoth JH, et al. The eﬀects of a unilateral ultrasound-guided renal

biopsy on renal function in healthy sedated cats. Vet Radiol Ultrasound 2000;41:57–62.

[12] Parrish AE. Complications of percutaneous renal biopsy: a review of 37 years’ experience.

Clin Nephrol 1992;38:135–41.

[13] McGill DB, Rakela J, Zinsmeister AR, Ott BJ. A 21-year experience with major

hemorrhage after percutaneous liver biopsy. Gastroenterology 1990;99:1396–400.

[14] McVay PA, Toy PTCY. Lack of increased bleeding after liver biopsy in patients with mild

hemostatic abnormalities. Am J Clin Pathol 1990;94:747–53.

[15] Bigge LA, Brown DJ, Pennick DG. Correlation between coagulation proﬁle ﬁndings and

bleeding complications after ultrasound-guided biopsies: 434 cases (1993–1996). J Am
Anim Hosp Assoc 2001;37:228–33.

[16] Mezzano D, Tagle R, Pais E, et al. Endothelial cell markers in chronic uremia: relationship

with hemostatic defects and severity of renal failure. Thromb Res 1997;88:465–72.

[17] Vaden SL. Renal biopsy. How and why. In: Proceedings of the 18th Annual Veterinary

Medical Forum, Seattle, 2000. p. 675.

[18] Larrain C, Langdell TD. The hemostatic defect of uremia. II. Investigation of dogs with

experimentally produced acute urinary retention. Blood 1956;11:1067–72.

[19] Osborne CA, Low DG, Jessen CR. Renal parenchymal response to needle biopsy. Invest

Urol 1972;9:463–9.

[20] Nash AS, Boyd JS, Minto W, Wright NG. Renal biopsy in the normal cat: an examination

of the eﬀects of a single biopsy. Res Vet Sci 1983;34:347–56.

[21] Stone EA, Barsanti JA. Diagnostic tests. In: Urologic surgery of the dog and cat.

Philadelphia: Lea & Febiger; 1992. p. 37–52.

[22] Osborne CA, Bartges JW, Polzin DJ, et al. Percutaneous needle biopsy of the kidney.

Indications, applications, technique and complications. Vet Clin North Am Small Anim
Pract 1996;26:1461–504.

[23] Patterson JM. Laparoscopy in small animal medicine. In: Kirk RW, editor. Current

veterinary therapy VII: small animal practice. Philadelphia: WB Saunders; 1980. p. 969–73.

[24] Stone EA, Barsanti JA. General surgical approaches. In: Urologic surgery of the dog and

cat. Philadelphia: Lea & Febiger; 1992. p. 100–6.

[25] Sweet EI, Davidson AJ, Hayslett JP. Complications of needle biopsy of the kidney in the

dog. Radiology 1969;92:849–54.

[26] Nash AS, Boyd JS, Minto AW, Wright NG. Renal biopsy in the normal cat: an

examination of the eﬀects of repeated needle biopsy. Res Vet Sci 1986;40:112–7.

[27] Jennette JC, et al. Heptinstall’s pathology of the kidney. 5th edition. Philadelphia:

Lippincott-Raven; 1998.

[28] Greenberg A, et al. Primer on kidney diseases. 3rd edition. San Diego: Academic Press;

2001.

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S.L. Vaden / Vet Clin Small Anim 34 (2004) 887–908

New and unusual causes of acute

renal failure in dogs and cats

Jennifer E. Stokes, DVM

S. Dru Forrester, DVM, MS

Department of Small Animal Clinical Sciences, C247 Veterinary Teaching Hospital,

College of Veterinary Medicine, University of Tennessee, 2407 River Drive,

Knoxville, TN 37996, USA

College of Veterinary Medicine, Western University of Health Sciences,

309 East 2nd Street, College Plaza, Pomona, CA 91766–1854, USA

This article was written with the intent to provide a source for easy

reference, summarizing in one location newly recognized and unusual causes
of acute renal failure (ARF) in dogs and cats. Several of the causes discussed
in this article have been described in other sources

[1–7]

. New or unusual

causes of ARF in dogs and cats include infectious diseases (leptospirosis,
borreliosis, and babesiosis), nephrotoxicants (aminoglycosides, vitamin D,
nonsteroidal anti-inﬂammatory drugs [NSAIDs]). and plant material (lilies
and raisins/grapes).

Plant toxicity

Development of ARF in cats caused by lily ingestion has been recognized

since at least 1989

. Members of the genus Lilium (Easter lily, tiger lily,

stargazer lily, and Asiatic hybrid lily) are potentially nephrotoxic to cats,
a species that seems uniquely aﬀected

. Nephrotoxicity has also been

attributed to members of the genus Hemerocallis (common daylily), which
are not true lilies but bear a lily-shaped ﬂower and are a perennial native to
eastern Asia that bloom throughout late spring to early autumn

Ingestion of ﬂowers or leaves of these plants can cause intoxication, but
the toxic principle and dose are unknown

[9,10]

. Toxicity in an individual

may be aﬀected by host factors, including gastrointestinal absorption and
preexisting renal disease, as well as by plant factors, such as stage, season,

* Corresponding author.
E-mail address:

jstokes4@utk.edu

(J.E. Stokes).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.006

Vet Clin Small Anim

34 (2004) 909–922

and potential toxic variability in diﬀerent parts of the plant

. Two other

plants commonly referred to as lilies, the calla lily and peace lily, are not
nephrotoxic as outlined in

Box 1 [9]

Lily intoxication causes gastrointestinal signs (vomiting and ptyalism),

neurologic signs (ataxia, depression, tremors, and seizures), and ARF

[9,10]

Gastrointestinal abnormalities and lethargy usually occur within hours of
plant ingestion

[8,9]

. ARF typically develops within 5 days after ingestion,

and cats may become oliguric or anuric

[9–11]

. Renomegaly and renal pain

have been described. Cats diagnosed with ARF have severe azotemia. In one
report of six cats with Easter lily–induced ARF, mean blood urea nitrogen
(BUN) was 215 mg/dL (range: 15–34 mg/dL) and mean serum creatinine
was 22.3 mg/dL (range: 0.8–2.3 mg/dL)

. Evidence of tubular damage

may be apparent on urinalysis, and aﬀected cats have been documented to
have cylindruria, glucosuria, and proteinuria

[8,10,11]

Successful treatment of lily toxicity includes early recognition, gastroin-

testinal decontamination, and ﬂuid diuresis. Cats that develop ARF may
require dialysis as part of their management, and the mortality rate for cats
with ARF ranges from 50% to 100%

[8–10]

. Cats that are decontaminated

within 6 hours of ingestion may not develop ARF, whereas those treated
more than 18 hours after exposure typically do so

[8,10,11]

. Thus, a guarded

prognosis is warranted for cats with ARF, whereas those with just
gastrointestinal disease usually do well. Chronic renal failure is a possible
sequela in cats surviving acute stages of renal failure

There are no histopathologic reports of daylily toxicity cases, but

histopathologic changes in the kidneys of cats that have ingested Lilium
sp include acute renal tubular necrosis, polarized crystals in the collecting
tubules, and renal tubular epithelial cell regeneration

[8,10]

. The toxic agent

causes an acute regional destruction of renal tubular epithelial cells;
basement membranes are usually intact, allowing possible regeneration

Box 1. Toxic and nontoxic plants known commonly as lilies

Toxic lilies

Calla lily (Zanteddeschia spp)
Daylily (Hemerocallis dumortieri, H fulvi, H graminea,

H seiboldi)

Easter lily (Lilium longiflorum)
Japanese show lily (L hybidium)

Nontoxic lilies

Peace lily (Spathiphyllum spp)
Rubrum lily (L rubrum)

Tiger lily (L tigrinum)

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Raisin and grape toxicity

Since 2001, there have been three published reports of acute gastrointes-

tinal and renal toxicity developing in 20 dogs after ingestion of large quantities
of raisins or grapes. Most of the cases have occurred since 1998 in the United
States and United Kingdom

[12–14]

. Aﬀected dogs had consumed fresh or

commercial grapes, both red and white varieties, as well as organic products,
grape crushings, or fermented grapes from wineries. Raisins implicated in
aﬀected dogs were usually commercial sun-dried raisins of diﬀerent brands

. The estimated amount of ingested raisins or grapes was known for 7 dogs

and ranged from 0.41 to 1.9 oz/kg of body weight

[12,13]

. To date, a speciﬁc

toxin has not been identiﬁed and screening results for contaminants, including
mycotoxins and heavy metals, have been negative

. There are no known

reports of raisin or grape toxicity in other species.

Vomiting and lethargy develop within hours of raisin or grape ingestion,

and anorexia, diarrhea, lethargy, and abdominal pain have also been
reported

. Acute renal failure developed within 24 to 72 hours, and

many dogs became oliguric or anuric. Fatality in dogs with ARF has ranged
from 50% to 75%, and oliguria or anuria was associated with a poor
prognosis

. All dogs that have recovered have been managed aggres-

sively, including use of peritoneal dialysis

. Histopathologic ﬁndings

include renal tubular necrosis, metastatic mineralization of numerous
tissues, and evidence of renal tubular epithelial regeneration

[12,14]

. Dogs

that ingest large quantities of grapes or raisins should be treated aggressively
with decontamination of the gastrointestinal tract, followed by intravenous
diuresis and close monitoring of renal function, including urine output

Vitamin D toxicosis

Vitamin D intoxication is a well-known cause of illness in dogs and cats.

Historically, the most common sources of vitamin D ingestion are
rodenticide products. Rodenticides are second only to insecticides in
prevalence of pesticide exposure

[2,15]

. Cholecalciferol (vitamin D

) is sold

over the counter under a variety of trade names and is formulated as
granules, ﬂakes, cakes and briquettes, typically as a 0.075% product

. The

major pathophysiologic eﬀect of vitamin D intoxication is dramatic
hypercalcemia, which can cause ARF. The toxic dose in dogs has been
reported as 1.5 to 8 mg/kg, but the toxic dose for cats is unknown

. In

general, puppies are more susceptible to vitamin D toxicosis than adult dogs
and cats are more sensitive than dogs

Toxicosis can also result from accidental ingestion of medications

containing vitamin D that are used for treatment of hypophosphatemic
disorders, hypoparathyroidism, osteomalacia, osteoporosis, and renal
failure in people

. A recently described vitamin D toxicant is calcipo-

triene, also known as calcipotriol, which is a synthetic analogue of calcitriol.

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J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

Calcipotriene is the active ingredient in Dovonex, a topical medication used
to treat people with psoriasis

. Ingestion of calcipotriene has been

associated with severe hypercalcemia, ARF, soft tissue mineralization, and
death in dogs

[2,3,16]

. There is little pharmacokinetic or toxicity informa-

tion in people or animals. An oral dose of 3.6 lg/kg/d for 7 days resulted in
renal failure in dogs, whereas a single dose of 37 lg/kg of body weight is
toxic in dogs

. Within 12 to 24 hours after ingestion, dogs exhibit

vomiting, depression, anorexia, polyuria, and diarrhea

. Hypercalcemia,

hyperphosphatemia, and hypercalcemic nephropathy can occur within 18 to
72 hours

Treatment of vitamin D toxicosis, regardless of the speciﬁc cause, is

focused on managing ARF and hypercalcemia. Saline diuresis, furosemide,
and glucocorticoids are standard therapies. Calcitonin and pamidronate
sodium, a second-generation bisphosphonate, have also been used to
manage hypercalcemia

[15–18]

. The two medications used together are less

eﬀective than either alone

. Pamidronate (1.3–2 mg/kg administered

intravenously) has been reported to reverse cholecalciferol-induced hyper-
calcemia in research dogs and prevented or reversed tubular necrosis and
mineralization caused by vitamin D toxicosis

. Prognosis after vitamin D

toxicosis is guarded, especially when intoxication is associated with a calcium
X phosphorus product of greater than 60 mg

/dL

. Death can occur weeks

after ingestion as a sequela from calciﬁcation of cardiac tissue

[3,16,19]

Antibiotics

Systemic use of aminoglycoside antibiotics has been known to cause

nephrotoxicity in animals since the 1970s

[20–22]

. Few clinical case reports

describing aminoglycoside-induced ARF have been published since the
1980s. A decrease in the number of reported intoxications may be partially
the result of an increased awareness of the toxic potential of aminoglyco-
sides, changes in dosing frequency, monitoring of drug serum concentra-
tions, and availability of safer antimicrobials with a similar spectrum of
action. Two unusual cases of aminoglycoside-induced ARF in cats have
been reported recently

[23,24]

A case of fatal nephrotoxicosis in a cat was associated with topical

administration of gentamicin to a soft tissue wound. A dose of 500 mg was
applied twice to an open wound during lavage. Serum gentamicin concen-
tration measured approximately 8 hours after the cat received topical
treatment was more than ﬁve times greater than therapeutic concentrations.
The cat was diagnosed with oliguric ARF within 4 days of receiving topical
gentamicin

Another report describes paromomycin-induced ARF in four cats

Paromomycin is an aminoglycoside that is poorly absorbed from the
gastrointestinal tract, and as little as 3% may appear in urine after oral

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J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

administration. All cats in the report were being treated for conﬁrmed or
suspected enteric trichomoniasis or cryptosporidiosis. There has been
increasing recognition of cryptosporidiosis and trichomoniasis as causes
of diarrhea in cats, and cats may serve as a zoonotic host for people.
Paromomycin has been used to treat resistant trichomoniasis infections in
people successfully. Two 1994 publications describe the use of paromomycin
(125–165 mg/kg administered orally every 12 hours for 5 days) as a safe and
eﬀective treatment for cryptosporidiosis in a small number of cats

[25,26]

. In

the four cats that developed renal failure, the paromomycin doses ranged
from 70 to 208 mg/kg administered orally every 12 hours for less than 5 days

. All four cats developed lethargy, vomiting, anorexia, dehydration, and

azotemia within 3 to 4 days of initiating therapy

. Cats were hospitalized

for 4 to 5 days and clinically improved with ﬂuid therapy. All cats had
improvement or resolution of azotemia within 6 to 11 months

. Three

cats also developed cataracts, and one was diagnosed with deafness after
treatment. Toxicity may have been caused in part by increased systemic
absorption of paromomycin from diseased bowel

Nonsteroidal anti-inﬂammatory drug and selective cyclooxygenase 2
inhibitor toxicity

NSAIDs and selective cyclooxygenase (COX) inhibitors are commonly

used in veterinary medicine for management of osteoarthritis and perioper-
ative pain

. Toxicity typically occurs as the result of an accidental

overdose but can develop with therapeutic doses in patients with concurrent
conditions predisposing to renal damage as listed in

Box 2 [28]

. Toxicity

caused by NSAIDs and selective COX inhibitors usually aﬀects the
gastrointestinal tract or kidneys, although hepatotoxicity has also been
associated with carprofen administration

All NSAID administration poses some risk for renal dysfunction,

although not equally

. Selective COX-2 inhibitors are reportedly safer

than nonselective inhibitors in people, causing fewer gastrointestinal and
renal side eﬀects than NSAIDs. Both classes of medication are potentially
toxic in dogs and cats, particularly because of species diﬀerences in the
expression and function of COX isoforms.

Mechanism of action

There are two isoforms of COX. COX-1 is primarily expressed

constitutively and is involved in the production of prostaglandins that
modulate normal physiologic functions in several organ systems, including
the kidneys, gastrointestinal tract, and platelets

. COX-2 expression is

inducible by bacterial endotoxins, cytokines, and growth factors and is
involved in the production of prostaglandins that modulate physiologic

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events in development, cell growth, and inﬂammation

. It has been

suggested that inhibitors of COX-2 decrease inﬂammation, whereas in-
hibition of COX-1 is responsible for side eﬀects associated with NSAID use,
including gastrointestinal ulceration and renal dysfunction

. COX-2 is

active not only during inﬂammation but has basal activity in the kidney and
brain in the absence of inﬂammatory stimulation

. Expression of COX-2

occurs at low levels in normal dogs but is greatly increased in dogs that are
volume depleted. Thus, administration of NSAIDs that preferentially
inhibit COX-2 may not spare patients from renal side eﬀects

Dogs are probably more susceptible to renal damage and ARF caused by

NSAIDs or COX-2 inhibitors than are people because of diﬀerences in renal
anatomy, distribution of COX-2 in renal tissues, and expression of COX-2
in volume-depleted states

[31,32]

. Species with unipapillary kidneys, such as

dogs, are more sensitive to NSAID-induced renal toxicity than are people,
who are a multipapillary species. There are also species diﬀerences in renal
medullary distribution of COX-1 and COX-2

. The papillary tip has

comparatively low renal oxygen tension, so it is more susceptible to injury
than other parts of the kidney. Dogs exhibit one or both COX isoforms in
the papillary interstitial cells, which are a rich source of prostaglandins

Interstitial cells are the ﬁrst cell type to undergo renal papillary necrosis
caused by NSAIDs and are probably the most susceptible cell in the renal
papilla because of their expression of COX isoforms

. There are

signiﬁcant interspecies diﬀerences in COX-2 localization, with wide distri-
bution of COX-2 in dogs. There is also a diﬀerence in the eﬀects of

Box 2. Conditions predisposing to acute renal failure

Nephrotoxins
Preexisting renal disease
Old age
Cardiovascular disease
Hypoalbuminemia
Liver disease
Acidosis
Diabetes mellitus
Electrolyte imbalances
Hypoadrenocorticism
Hypotension
Hyperviscosity syndrome
Hypovolemia
Fever
Dehydration
Sepsis

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hypovolemia on COX-2 activity. In people, sodium depletion does not
signiﬁcantly alter COX-2 activity. Kidneys from sodium-depleted dogs show
an approximately threefold increase in COX-2 immunoreactivity in the
macula densa and thick ascending limb of the loop of Henle

Constitutive expression of COX-2 at several sites in dogs indicates that this
isoform is involved in regulation of normal renal functions in these species,
as is also indicated by its upregulation in volume-depleted dogs

. COX-2

may play an important role in maintaining renal blood ﬂow in volume-
depleted dogs

Toxic principles

The prevalence of NSAID-induced renal insuﬃciency in companion

animals is unknown

. Although ARF in dogs has been associated with

administration of phenylbutazone, aspirin, ibuprofen, carprofen, naproxen,
and ﬂunixin meglumine, ibuprofen has become the most common generic
drug generating calls to the National Animal Poison Control Center for
dogs and cats

[28,33]

. The overall occurrence of ARF caused by NSAIDs is

low. Most dogs that develop ARF with NSAID treatment either ingest
excess amounts of the drug or have a concurrent disorder increasing their
susceptibility to ARF

Only ibuprofen-induced ARF in dogs has been well described in the

literature

. In one review, ARF subsequent to ibuprofen was more likely

to develop when it took longer for the patient to receive medical
intervention

. Appropriate early intervention, including gastrointestinal

decontamination, was more eﬀective for managing gastrointestinal disease
than for preventing renal disease

. ARF generally occurred 36 hours or

longer after exposure, whereas gastrointestinal signs occurred within a few
hours

. In experiments, a dose of 300 mg/kg was required to produce

ARF in dogs. In this study, 32 cases of ARF had exposure doses of less than
300 mg/kg

. Another source reports that ARF caused by ischemia is

likely at doses of greater than or equal to 175 to 200 mg/kg in dogs

. Cats

are susceptible to ibuprofen doses approximately one half of those reported
toxic in dogs, although no experimental data are available to conﬁrm and
predict toxicosis in this species

Volume depletion increases the susceptibility to the nephrotoxic eﬀects

of NSAIDs and COX-2 inhibitors in dogs

. Volume depletion leads to

upregulation of COX-2 protein in the macula densa in dogs

. Canine

studies show that when the renin-angiotensin system is activated,
NSAIDs decrease renal blood ﬂow and glomerular ﬁltration rate
(GFR)

. Many NSAIDs have been shown to decrease renal function

or renal blood ﬂow. Normal healthy dogs given carprofen or ketoprofen
had a signiﬁcant decrease in serum creatinine clearance for the 24-hour
period from 24 to 48 hours after drug administration compared with
control dogs

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J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

In volume-depleted dogs, intravenous celecoxib reduced urine output by

57%, sodium excretion by 70%, renal plasma ﬂow by 65%, and GFR by
58%

. The same values were decreased by 57%, 33%, 33%, and 27%,

respectively, when indomethacin was administered

. Indomethacin (10

mg/kg) did not alter renal blood ﬂow in awake dogs but was associated with
a 40% decrease in renal blood ﬂow in anesthetized dogs undergoing surgery

. Anesthetized dogs given intravenous ibuprofen (10 mg/kg) or in-

domethacin (2.5 mg/kg) exhibited a 62% increase in renal vascular
resistance

. In one additional study, an experimental COX-2 inhibitor

did not decrease renal arteriolar blood ﬂow in volume-depleted dogs,
whereas indomethacin did

[37]

Clinical presentation and management

A tentative diagnosis of NSAID-induced ARF is usually made based on

history, physical examination, and results of laboratory evaluation. Most
dogs have an acute onset of clinical signs

. Inappetence, vomiting,

diarrhea, and melena are common and may result from gastrointestinal
ulceration or ARF. Signs of gastrointestinal ulceration are common and
typically precede ARF

. There is no speciﬁc therapy for NSAID-induced

ARF, just supportive care. When dogs ingest potentially toxic doses,
management with gastric decontamination and ﬂuid diuresis (to prevent
decreased renal blood ﬂow and to counteract hypotension) is recommended

. Administration of sodium bicarbonate can be added to enhance

ibuprofen excretion

. The use of NSAIDs in patients at higher risk (ie,

patients with cardiovascular disease) should be avoided if possible

Most dogs that develop NSAID-induced ARF have a favorable prognosis

. They generally respond well after appropriate treatment for 5 to 10

days

Emerging and re-emerging infectious diseases

Babesiosis

Babesiosis is an emerging infectious disease in dogs in the United States.

Currently, there are two distinct species known to cause disease in dogs:
Babesia canis

and B gibsoni. Genetic testing has identiﬁed three types of

B canis

, which are larger organisms than B gibsoni. These three subspecies,

B canis canis

, B canis vogeli, and B canis rossi, may be classiﬁed into separate

species

. B canis canis is endemic in the southeastern United States and is

common in Greyhounds. A 1992 study found that 46% of Florida racing
Greyhounds had positive titers for B canis

. Most adults are subclinical

carriers, but the disease may cause severe anemia and ‘‘fading puppy’’
syndrome in neonates. Adults may also develop severe hemolytic anemia
and thrombocytopenia. B gibsoni was ﬁrst reported in the United States in

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J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

1968, but this dog was shipped from Malaysia and most likely had been
infected before its arrival in the United States

. In 1991, 11 dogs from

California were diagnosed with B gibsoni infection, and these dogs de-
veloped severe hemolytic anemia and thrombocytopenia

. Since that

time, B gibsoni infection has been reported in eight other states, and almost
all cases have involved American Pit Bull Terriers or American Staﬀordshire
Terriers

. Variability in pathogenicity of B gibsoni isolates has been

described, resulting in diﬀerences in clinical syndromes in diﬀerent geo-
graphic locations.

Babesiosis rarely causes ARF in dogs, and one report found an incidence

of 2.2% in 134 dogs

. Possible causes for renal failure in dogs with

babesiosis include anemic hypoxia, hypovolemia, hemoglobinuric nephrop-
athy, and myoglobinuric nephropathy secondary to rhabdomyolysis

[39,40]

Many more dogs with babesiosis have hemoglobinuria than develop ARF
or renal lesions, indicating that multiple factors may lead to renal
dysfunction and not hemoglobinuria alone. Anoxia, renal blood ﬂow
reduction, hypotension, and renal ischemia probably play a more important
role in the development of ARF than does hemoglobinuria. Hypoxia results
in more renal tubular injury than does hemoglobin, and nephrotoxic eﬀects
of hemoglobin are highly individual

Borreliosis

Lyme disease is caused by the spirochete bacterium Borrelia burgdorferi

and is transmitted by ticks of the genus Ixodes. Transmission requires 48
hours of tick attachment

[41]

. Borreliosis was ﬁrst documented in people in

the United States in 1969, and canine borreliosis was ﬁrst described in 1984

. Seroprevalence in people and dogs varies a great deal based on

geographic location. Most infected dogs do not have clinical signs. Canine
borreliosis usually manifests as lameness, fever, lethargy, and anorexia;
cardiac and neurologic signs are rare

. Dogs respond quickly to treatment,

with clinical improvement typically seen within 48 hours of initiating
appropriate antimicrobial therapy

Although the incidence is unknown, Lyme disease has been associated

with a severe protein-losing glomerulopathy and ARF in dogs, but no
animal model exists to study the disease

. ‘‘Lyme nephritis,’’ as it has been

described, was reported in 49 dogs between 1987 and 1992 in areas endemic
for Lyme disease

. Cases were diagnosed in early spring through late

autumn, which is the time when most cases of Lyme disease are diagnosed in
dogs and people. The mean age of aﬀected dogs was 5.6 years, which is
younger than dogs with other glomerulopathies (mean age of 7.1 years).
Labrador Retrievers, Golden Retrievers, and Shetland Sheepdogs were
breeds identiﬁed as being predisposed to developing Lyme nephritis

[6,42]

The disease progressed much more quickly than is typical of other
glomerulopathies, which are chronic in nature. Aﬀected dogs developed

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J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

an acute progressive renal failure associated with azotemia, uremia, pro-
teinuria, peripheral edema, and body cavity eﬀusion

[41]

. Hypertension was

also present in many cases

. The duration of clinical illness ranged from 24

hours to 8 weeks, and most dogs presented with a sudden onset of anorexia,
vomiting, lethargy, and weight loss. Neurologic signs were noted in about
15% of dogs. Hematologic abnormalities included nonregenerative anemia,
thrombocytopenia, and stress leukogram

. Abnormalities on urinalysis

included proteinuria, hemoglobinuria, glucosuria, and cylindruria with or
without hematuria. Clinical disease progressed rapidly in most dogs, leading
to euthanasia or death within 1 to 2 weeks, although 3 dogs lived for several
months

Lyme nephritis seems to be a unique combination of immune-mediated

glomerulonephritis, diﬀuse tubular necrosis with regeneration, and lympho-
cytic-plasmacytic interstitial nephritis

. Of the 49 cases described, immune-

mediated membranoproliferative glomerulonephritis was found in 43 cases,
membranous glomerulonephritis in 5 cases, and amyloidosis in 1 case

Subendothelial deposits of IgG, IgM, and C3 were present along the
glomerular membrane

. Of the 18 dogs tested, all were serologically

positive for B burgdorferi

. Other dogs had spirochetes in the renal tissue,

and in 1 case, a urine culture was positive for B burgdorferi

. Approximately

30% of dogs had a history of lameness, usually within 3 to 6 weeks of
presentation, and some had been treated for Lyme disease with clinical
improvement

. Almost 30% had been vaccinated for Lyme disease

Further research needs to be done to determine the role, if any, of vaccination
in the development of acute, progressive, fatal renal lesions that occur in
canine Lyme nephritis

[41]

. The underlying cause has yet to be discovered and

may be linked to B burgdorferi infection or to an unrelated agent.

Leptospirosis

Leptospirosis is a worldwide zoonotic disease that aﬀects people, dogs,

livestock, and many other domestic and wild species. Leptospirosis is caused
by a spirochete bacterium and has been reported in dogs in the United
States and Canada for more than 100 years

. There are greater than 220

pathogenic species and many more that are avirulent

[44]

. Pathogenic

serovars that infect dogs include canicola, icterohaemorrhagiae, bratislava,
grippotyphosa, hardjo, pomona, australis, ballum, and autumnalis and are
of the genus Leptospira and the species L interrogans or L kirschneri

Each serovar is maintained in the environment by one or more reservoir
hosts, which is a species that typically remains asymptomatic when infected
but sheds large numbers of infective organisms into the environment (

Table

). Dogs are a reservoir host for the serovar canicola but also serve as

secondary hosts for other serovars that cause illness. Leptospiral infection
can be asymptomatic or cause a clinical syndrome of ARF, hepatic disease,
coagulation disorders, or a combination of syndromes.

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Between 1970 and 1982, the prevalence of canine leptospirosis decreased

in the United States and Canada, most likely because of the widespread use
of a bivalent vaccine that became available in the early 1970s

. The

bivalent vaccine prevents disease caused by the serovars canicola and
icterohaemorrhagiae but not by other serovars, because protective anti-
bodies are serovar speciﬁc. In contrast, the prevalence of leptospirosis
increased in dogs in the United States and Canada between 1983 and 1998,
with an annual increase of 1.2 cases per 100,000 dogs examined

Historically, most cases of canine leptospirosis were caused by the serovars
canicola and icterohaemorrhagiae, but recent clinical reports have shown
that the disease is now most commonly caused by the serovars grippoty-
phosa, pomona, and bratislava

[45–48]

. The change in the epidemiology of

canine leptospirosis may be the result of urbanization leading to increased
direct or indirect exposure of dogs to wildlife and livestock reservoir hosts as
well as the result of a marked decrease in disease caused by the serovars
canicola and icterohaemorrhagiae.

A retrospective review of leptospirosis in 677 dogs found that male dogs 4

years of age or older and herding dogs, hounds, and working dogs were at
a signiﬁcantly higher risk of disease

. Despite these risk factors,

leptospirosis is diagnosed in toy breeds and young dogs

[47,50]

. There has

also been an association between increased rainfall and increased number of
cases of canine leptospirosis

. Most clinical cases of leptospirosis

reported in dogs in the past two decades have been of ARF, with or
without hepatic involvement. Few cases have manifested as hepatic disease
alone. With early recognition, appropriate antibiotic therapy, aggressive
ﬂuid therapy, and sometimes dialysis, many dogs survive infection

[52]

. A

mortality rate of 11% to 33% as been reported

[45,47,53]

. Based on clinical

reports, 33% to 40% of dogs develop chronic renal failure as a sequela to
the acute infection

[46–48,54]

Miscellaneous causes

Isolated case reports have also described ARF in dogs as the result of

a variety of causes. In 1995, acute intrinsic renal failure and coagulopathy
secondary to a snake bite (Vipera aspis) was reported in a dog

. The dog

Table 1
Leptospiral serovars and their primary (reservoir) host(s)

Species

Serovar

Primary (reservoir) hosts

Leptospira interrogans

Bratislava

Rat, pig

Canicola

Dog

Hardjo

Cow

Icterohaemorrhagiae

Rat

Pomona

Cow, pig, skunk, opossum

L kirshneri

Grippotyphosa

Raccoon, skunk, vole, opossum

919

J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

developed acute tubular necrosis, glomerular hypercellularity, mesangial
lysis, and cylindruria. Snakebites are a source of nephrotoxins that cause
renal ischemia, alterations in renal perfusion, and coagulopathies. The toxic
eﬀect of a snake bite depends on the species of snake, time of year, weather,
and time between the bite and medical intervention

A 2002 report describes toxicity in a dog ingesting isoniazid, an

antimicrobial agent used to treat tuberculosis in people and animals

[56]

The dog developed severe seizures and presumptive exertional rhabdomyol-
ysis. Fatal anuric ARF with tubular necrosis developed, attributed to
rhabdomyolysis-induced myoglobinemia and myoglobinuria. Myoglobinu-
ria has been implicated in acute renal tubular necrosis

[56]

Summary

Despite advances in veterinary medicine, ARF continues to be

associated with high morbidity and mortality. The discovery of new
causes of ARF and the re-emergence of several infectious diseases that can
cause ARF necessitate being aware of changes in the ﬁeld of veterinary
medicine. Although it is important for veterinarians to be aware of
common causes of ARF in dogs and cats, emphasis should be placed on
recognizing predisposing factors and, when possible, correcting abnormal-
ities that may lead to ARF.

References

[1] Langston CE, Heuter KJ. Leptospirosis. A re-emerging zoonotic disease. Vet Clin North

Am Small Anim Pract 2003;33(4):791–807.

[2] Murphy MJ. Rodenticides. Vet Clin North Am Small Anim Pract 2002;32(2):469–84.
[3] Welch SL. Oral toxicity of topical preparations. Vet Clin North Am Small Anim Pract

2002;32(2):443–53.

[4] Albretsen JC. Oral medications. Vet Clin North Am Small Anim Pract 2002;32(2):421–42.
[5] Rosypal AC, Zajac AM, Lindsay DS. Canine visceral leishmaniasis and its emergence in

the United States. Vet Clin North Am Small Anim Pract 2003;33(4):921–37.

[6] Littman MP. Canine borreliosis. Vet Clin North Am Small Anim Pract 2003;33(4):827–62.
[7] Boozer AL, Macintire DK. Canine babesiosis. Vet Clin North Am Small Anim Pract 2003;

33(4):885–904.

[8] Brady MA, Janovitz EB. Nephrotoxicosis in a cat following ingestion of Asiatic hybrid lily

(Lilium sp). J Vet Diagn Invest 2000;12(6):566–8.

[9] Hadley RM, Richardson JA, Gwaltney-Brant SM. A retrospective study of daylily

toxicosis in cats. Vet Hum Toxicol 2003;45(1):38–9.

[10] Langston CE. Acute renal failure caused by lily ingestion in six cats. J Am Vet Med Assoc

2002;220(1):36, 49–52.

[11] Gulledge L, Boos D, Wachsstock R. Acute renal failure in a cat secondary to Tiger Lily

(Lilium tigrinum) toxicity. Feline Pract 1997;25:38–9.

[12] Gwaltney-Brant S, Holding JK, Donaldson CW, Eubig PA, Khan SA. Renal failure

associated with ingestion of grapes or raisins in dogs. J Am Vet Med Assoc 2001;218(10):
1555–6.

920

J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

[13] Campbell A, Bates N. Raisin poisoning in dogs. Vet Rec 2003;152(12):376.
[14] Penny D, Henderson SM, Brown PJ. Raisin poisoning in a dog. Vet Rec 2003;152(10):308.
[15] Rumbeiha WK, Fitzgerald SD, Kruger JM, et al. Use of pamidronate disodium to reduce

cholecalciferol-induced toxicosis in dogs. Am J Vet Res 2000;61(1):9–13.

[16] Fan TM, Simpson KW, Trasti S, Birnbaum N, Center SA, Yeager A. Calcipotriol toxicity

in a dog. J Small Anim Pract 1998;39(12):581–6.

[17] Fooshee SK, Forrester SD. Hypercalcemia secondary to cholecalciferol rodenticide

toxicosis in two dogs. J Am Vet Med Assoc 1990;196(8):1265–8.

[18] Dougherty SA, Center SA, Dzanis DA. Salmon calcitonin as adjunct treatment for vitamin

D toxicosis in a dog. J Am Vet Med Assoc 1990;196(8):1269–72.

[19] Volmer P. Oral toxicity of skin and eye products. Presented at the 18th Annual Veterinary

Medical Forum, American College of Veterinary Internal Medicine, Seattle, May 24–27, 2000.

[20] Schultze RG, Winters RE, Kauﬀman H. Possible nephrotoxicity of gentamicin. J Infect

Dis 1971;124(Suppl 124):145.

[21] Engle JE, Abt AB, Schneck DW, Schoolwerth AC. Netilmicin and tobramycin.

Comparison of nephrotoxicity in dogs. Invest Urol 1979;17(2):98–102.

[22] Reiner NE, Bloxham DD, Thompson WL. Nephrotoxicity of gentamicin and tobramycin

given once daily or continuously in dogs. J Antimicrob Chemother 1978;4(Suppl A):85–101.

[23] Mealey KL, Boothe DM. Nephrotoxicosis associated with topical administration of

gentamicin in a cat. J Am Vet Med Assoc 1994;204(12):1919–21.

[24] Gookin JL, Riviere JE, Gilger BC, Papich MG. Acute renal failure in four cats treated with

paromomycin. J Am Vet Med Assoc 1999;215(12):1806, 1821–3.

[25] Barr SC, Jamrosz GF, Hornbuckle WE, Bowman DD, Fayer R. Use of paromomycin for

treatment of cryptosporidiosis in a cat. J Am Vet Med Assoc 1994;205(12):1742–3.

[26] Barr SC, Guilford WG, Jamrosz GF, Hornbuckle WE, Bowman DD. Paromomycin for

the treatment of cryptosporidium in dogs and cats [abstract]. J Vet Intern Med 1994;8:177.

[27] Forsyth SF, Guilford WG, Pfeiﬀer DU. Eﬀect of NSAID administration on creatinine

clearance in healthy dogs undergoing anaesthesia and surgery. J Small Anim Pract 2000;
41(12):547–50.

[28] Forrester SD, Troy GC. Renal eﬀects of nonsteroidal antiinﬂammatory drugs. Compend

Contin Educ Pract Vet 1999;21(10):910–9.

[29] MacPhail CM, Lappin MR, Meyer DJ, Smith SG, Webster CR, Armstrong PJ.

Hepatocellular toxicosis associated with administration of carprofen in 21 dogs. J Am
Vet Med Assoc 1998;212(12):1895–901.

[30] Lobetti RG, Joubert KE. Eﬀect of administration of nonsteroidal anti-inﬂammatory drugs

before surgery on renal function in clinically normal dogs. Am J Vet Res 2000;61(12):1501–7.

[31] Khan KN, Venturini CM, Bunch RT, et al. Interspecies diﬀerences in renal localization of

cyclooxygenase isoforms: implications in nonsteroidal antiinﬂammatory drug-related
nephrotoxicity. Toxicol Pathol 1998;26(5):612–20.

[32] Silverman LR, Khan KN. ‘‘Have you seen this?’’ Nonsteroidal anti-inﬂammatory drug-

induced renal papillary necrosis in a dog. Toxicol Pathol 1999;27(2):244–5.

[33] Poortinga EW, Hungerford LL. A case-control study of acute ibuprofen toxicity in dogs.

Prev Vet Med 1998;35(2):115–24.

[34] Villar D, Buck WB, Gonzalez JM. Ibuprofen, aspirin and acetaminophen toxicosis and

treatment in dogs and cats. Vet Hum Toxicol 1998;40(3):156–62.

[35] Stubanus M, Riegger GA, Kammerl MC, Fischereder M, Kramer BK. Renal side-eﬀects

of cyclo-oxygenase-type-2 inhibitor use. Lancet 2000;355(9205):753.

[36] Venturini CM, Isakson P, Needleman P. Non-steroidal anti-inﬂammatory drug-induced

renal failure: a brief review of the role of cyclo-oxygenase isoforms. Curr Opin Nephrol
Hypertens 1998;7(1):79–82.

[37] Black SC, Brideau C, Cirino M, et al. Diﬀerential eﬀect of a selective cyclooxygenase-2

inhibitor versus indomethacin on renal blood ﬂow in conscious volume-depleted dogs.
J Cardiovasc Pharmacol 1998;32(5):686–94.

921

J.E. Stokes, S.D. Forrester / Vet Clin Small Anim 34 (2004) 909–922

[38] Macintire DK. Babesia gibsoni—an emerging disease. Presented at the 20th American

College of Veterinary Internal Medicine Forum. Dallas, May 29–June 1, 2002.

[39] Lobetti RG, Jacobson LS. Renal involvement in dogs with babesiosis. J S Afr Vet Assoc

2001;72(1):23–8.

[40] Jacobson LS, Lobetti RG. Rhabdomyolysis as a complication of canine babesiosis. J Small

Anim Pract 1996;37(6):286–91.

[41] Greene CE, Appel MJG, Straubinger RK. Lyme borreliosis. In: Greene CE, editor.

Infectious diseases of the dog and cat. Philadelphia: WB Saunders; 1998. p. 282–93.

[42] Dambach DM, Smith CA, Lewis RM, Van Winkle TJ. Morphologic, immunohistochem-

ical, and ultrastructural characterization of a distinctive renal lesion in dogs putatively
associated with Borrelia burgdorferi infection: 49 cases (1987–1992). Vet Pathol 1997;34(2):
85–96.

[43] Ward MP. Clustering of reported cases of leptospirosis among dogs in the United States

and Canada. Prev Vet Med 2002;56(3):215–26.

[44] Kalin M, Devaux C, DiFruscia R, Lemay S, Higgins R. Three cases of canine leptospirosis

in Quebec. Can Vet J 1999;40(3):187–91.

[45] Adin CA, Cowgill LD. Treatment and outcome of dogs with leptospirosis: 36 cases (1990–

1998). J Am Vet Med Assoc 2000;216(3):371–5.

[46] Harkin KR, Gartrell CL. Canine leptospirosis in New Jersey and Michigan: 17 cases

(1990–1995). J Am Anim Hosp Assoc 1996;32(6):495–501.

[47] Brown CA, Roberts AW, Miller MA, et al. Leptospira interrogans serovar grippotyphosa

infection in dogs. J Am Vet Med Assoc 1996;209(7):1265–7.

[48] Birnbaum N, Barr SC, Center SA, Schermerhorn T, Randolph JF, Simpson KW.

Naturally acquired leptospirosis in 36 dogs: serological and clinicopathological features.
J Small Anim Pract 1998;39(5):231–6.

[49] Ward MP, Glickman LT, Guptill LE. Prevalence of and risk factors for leptospirosis

among dogs in the United States and Canada: 677 cases (1970–1998). J Am Vet Med Assoc
2002;220(1):53–8.

[50] Hrinivich K, Prescott JF. Leptospirosis in 2 unrelated dogs. Can Vet J 1997;38(8):509–10.
[51] Ward MP. Seasonality of canine leptospirosis in the United States and Canada and its

association with rainfall. Prev Vet Med 2002;56(3):203–13.

[52] Greene CE, Miller MA, Brown CA. Leptospirosis. In: Greene CE, editor. Infectious

diseases of the dog and cat. Philadelphia: WB Saunders; 1998. p. 273–81.

[53] Forrester SD, McMillan NS, Ward DL. Retrospective evaluation of acute renal failure in

dogs. J Vet Intern Med 2002;16:354.

[54] Rentko VT, Clark N, Ross LA, Schelling SH. Canine leptospirosis. A retrospective study

of 17 cases. J Vet Intern Med 1992;6(4):235–44.

[55] Puig J, Vilafranca M, Font A, Closa J, Pumarola M, Mascort J. Acute intrinsic renal

failure and blood coagulation disorders after a snakebite in a dog. J Small Anim Pract
1995;36(7):333–6.

[56] Haburjak JJ, Spangler WL. Isoniazid-induced seizures with secondary rhabdomyolysis and

associated acute renal failure in a dog. J Small Anim Pract 2002;43(4):182–6.

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Diagnosis of urinary tract infections

Joseph W. Bartges, DVM, PhD

Department of Small Animal Clinical Sciences, C247 Veterinary Teaching Hospital,

College of Veterinary Medicine, The University of Tennessee,

Knoxville, TN 37996–4544, USA

A urinary tract infection (UTI) occurs when there is a breach (either

temporary or permanent) in host defense mechanisms and suﬃcient
numbers of a virulent microbe are allowed to adhere, multiply, and persist
in a portion of the urinary tract. UTIs typically involve bacteria; however,
fungi and viruses may infect the urinary tract. Infection may predominate at
a single site, such as the kidney (pyelonephritis), ureter (ureteritis), urinary
bladder (cystitis), urethra (urethritis), prostate (prostatitis), or vagina
(vaginitis), or at two or more of these sites. Because a UTI may involve
more than one location, it may be more relevant to identify the infection
anatomically, that is, upper urinary tract (kidneys and ureters) versus lower
urinary tract (bladder, urethra, and prostate or vagina). The infection may
or may not produce clinical signs.

Incidence of urinary tract infections

The reported incidence of bacterial UTIs in dogs and cats is variable.

Bacterial UTI is estimated to aﬀect 14% of all dogs during their lifetime;
female dogs are more often aﬀected than male dogs

[1–5]

. Bacterial UTI is

more common in older cats (>10 years) than in younger cats, and the
incidence increases with age

[6–8]

. Occurrence of fungal UTI is seems to be

uncommon in dogs or cats

[9–12]

. Incidence of viral UTI is unknown;

however, viruses have been implicated as a cause of lower urinary tract
disease in cats

[13–17]

The pathogenesis of UTI represents a balance between uropathogenic

infectious agents and host resistance. UTI is treated with antimicrobial
agents; however, status of host defense mechanisms is important in
development of UTI and in successful treatment and prevention.

E-mail address:

jbartges@utk.edu

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.001

Vet Clin Small Anim

34 (2004) 923–933

Deﬁnition of terms

Urinary tract infection

UTI is deﬁned as adherence, multiplication, and persistence of an

infectious agent in the urogenital system. This often involves an organism
that is present normally in the distal urogenital tract. Most commonly, UTI
involves a bacterial organism.

Microburia

Microburia refers to the presence of microbes, typically bacteria, in urine.

Bacteruria

Bacteruria refers to presence of bacteria in urine. Identiﬁcation of

bacteria in urine is not synonymous with UTI. Bacteria may represent
contamination of a urine sample, particularly if the sample is collected by
voiding or urethral catheterization. Bacteria from the distal urogenital tract
may appear in urine collected by these methods, or urine may be contami-
nated after collection (eg, during transfer to storage containers in the
laboratory). Signiﬁcant bacteruria is used to describe bacteruria associated
with UTI. High bacterial numbers in a properly collected and cultured urine
sample indicate bacterial UTI. Small numbers of bacteria obtained from
urine of untreated patients usually indicate contamination. Asymptomatic
bacteruria refers to signiﬁcant bacteruria that is not associated with clinical
signs of UTI. This often occurs in animals with compromised host defenses,
such as glucocorticoid excess, diabetes mellitus, or infection with feline
immunodeﬁciency virus.

Funguria

Funguria refers to presence of fungi in urine. Normally, fungi are not

present in the urogenital system; when they are identiﬁed, it represents an
infection whether clinical signs exist or not.

Pyuria

Pyuria refers to presence of white blood cells (WBCs) in urine. Signiﬁcant

pyuria is deﬁned as ﬁnding greater than 3 to 5 WBCs (ie, neutrophils) per
high-power ﬁeld during sediment examination of urine collected by cys-
tocentesis. Finding greater than 5 to 10 WBCs per high-power ﬁeld in urine
collected by urethral catheterization or voiding is considered signiﬁcant.

Inﬂammation versus infection

Pyuria indicates urinary tract inﬂammation and is not synonymous with

UTI. Many disease processes result in inﬂammation of the urinary system

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

characterized by hematuria, pyuria, or proteinuria. The presence of one or
more of these does not indicate the cause or location of the disease within
the urogenital system. To identify the cause of inﬂammation, additional
testing, such as diagnostic imaging, urine culture, renal function tests,
endoscopy, urodynamic studies, or biopsy, is required.

Clinical ﬁndings

Historical information and physical examination ﬁndings

Dogs and cats with UTI may or may not have clinical signs of urinary

tract disease. Clinical signs associated with UTI are variable and depend on
the interaction of (1) virulence and numbers of the uropathogen, (2)
presence or absence of predisposing causes, (3) the body’s compensatory
response to infection, (4) duration of infection, and (5) site(s) of infection
(

). Pollakiuria, stranguria, dysuria, and inappropriate urination may

be observed with lower UTI. Animals with upper UTI may exhibit pain
(localized to one or both kidneys), hematuria, septicemia, or renal failure
(with resultant clinical signs if both kidneys are infected). If UTI is
associated with a predisposing condition, such as diabetes mellitus, hyper-
adrenocorticism, or urinary bladder neoplasia, clinical signs associated with
the predisposing condition may be present. Female dogs with vulvar
abnormalities, perivulvar dermatitis, or vaginal stenosis may have increased
risk for UTI

[18]

. Male cats with perineal urethrostomies also have increased

risk for UTI

. Some animals may have infection of both the upper and

lower urinary tracts, especially if renal failure exists.

Laboratory results

Unless septicemia or renal failure is present, results of complete blood cell

counts are normal. If septicemia is present, leukocytosis and a left shift may
occur. Lower UTI does not cause changes in results of routine laboratory
tests unless another disease process is present. With upper UTI, results of
serum biochemical analysis may be normal or may indicate renal failure (if
both kidneys are diseased). If UTI is associated with another disease,
changes in laboratory parameters may reﬂect the associated condition. In
cats, feline leukemia virus and feline immunodeﬁciency virus infection
increase risk of UTI

Results of diagnostic imaging studies and endoscopy

In many dogs and cats with UTI, results of diagnostic imaging studies are

normal. Survey abdominal radiographs may reveal uroliths, renomegaly, or
small kidneys or other defects that predispose to development of UTI. Some
dogs may have pelvic displacement of the urinary bladder (so-called ‘‘pelvic
bladder’’), which can be associated with urinary incontinence and UTI,

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

Table 1
Abnormalities that help to localize urinary tract infections

Sites of UTI

History

Physical examination

Laboratory

Diagnostic imaging studies

Lower

urinary
tract

Dysuria, pollakiuria
Urge to incontinence
Signs of abnormal detrusor

reﬂex (overﬂow incontinence,
large residual volume)

Gross hematuria at end of

micturition

Cloudy urine with abnormal

odor

No systemic signs
Recent catheterization or

urethrostomy

Small, painful, or thickened

urinary bladder

Palpable masses in urethra or

bladder

Flaccid bladder wall, large

residual volume

Abnormal micturition reﬂex
palpation of urethroliths

CBC: normal
Urinalysis: pyuria, hematuria,

proteinuria, bacteruria

Urine culture: signiﬁcant

bacteruria

Normal kidneys
Structural abnormalities of

lower urinary tract

urocystoliths and/or

urethroliths

thickening of bladder wall

and irregularity of mucosa

Rarely intraluminal gas

formation (emphysematous
cystitis)

Upper

urinary
tract

Polyuria, polydipsia
signs of systemic infection
renal failure

no detectable abnormalities
fever and other signs of

systemic infection

abdominal (renal) pain with

kidney(s) normal or enlarged

CBC:

leukocytosis

Urinalysis: pyuria, hematuria,

proteinuria, bacteruria, white
blood cell or granular casts

Imparied urine concentration
azotemia and other ﬁndings

of renal failure

Renomegaly
abnormal kidney shape
nephroliths, ureteroliths
dilated renal pelves, dilated

pelvic diverticula

evidence of outﬂow

obstruction

Acute

prostatitis
or prostatic
abscessation

Urethral discharge independent

of micturition

Signs of systemic infection
reluctance to urinate or

defecate

fever and other signs of

systemic infection

painful prostate and/or

painful abdomen

prostatomegaly or

asymmetric prostate

CBC:

leukocytosis

Urinalysis: pyuria, hematuria,

proteinuria, bacteruria

Cytologic evaluation of

prostate: inﬂammation or
infection

indistinct cranial border of

prostate

prostatomegaly
prostatic cysts
reﬂux of contrast medium

into prostate

Chronic

prostatitis

Recurrent UTI
Urethral discharge independent

of urination

dysuria

Often no detectable

abnormalities

prostatomegaly or

asymmetric prostate

CBC: normal
Urinalysis: pyuria,

hematuria, proteinuria,

bacteruria

prostatomegaly
prostatic cysts
prostatic mineralization

Abbreviations:

CBC indicates complete blood cell count; UTI, urinary tract infection.

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34
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923–933

although it is observed to occur in dogs without disease

. If no

abnormalities are found by survey abdominal radiography, ultrasonography
or contrast radiography should be performed. The upper urinary tract may
be evaluated by excretory urography, whereas the lower urinary tract may
be evaluated by contrast cystourethrography, double-contrast cystography,
and contrast vaginourethrography. A potential complication of performing
contrast radiography of the lower urinary tract is inducing an UTI.
Ultrasonography is a noninvasive technique and is useful in evaluating
echotexture and architecture of the urinary tract, except for the distal
urethra. In human beings, nuclear scintigraphy using technetium Tc 99m–
labeled dimercaptosuccinic acid (DMSA) may be useful for evaluating
pyelonephritis, although this technique has not been used in dogs

Endoscopy of the lower urinary tract may be useful in identifying

mucosal and intraluminal lesions that may predispose to UTI. In one study,
a urolith not visible by survey radiography was visualized by cystoscopy in
a cat

. Disadvantages of cystourethroscopy include anesthesia necessary

to perform the procedure, contamination and trauma to the lower
urogenital tract, and diﬃculty in performing the procedure in male cats.

Diagnosis

Urinalysis

A urinalysis should be performed routinely as part of a minimum

database. A complete urinalysis includes determining urine speciﬁc gravity
(USPG) using a refractometer, chemical analysis using analytic test pads on
dipsticks, and microscopic examination of urine sediment. Collection of
urine by cystocentesis is the preferred method when evaluating patients for
UTI. If infectious prostatitis or vaginitis is suspected, diﬀerent techniques
are indicated.

With UTI, the USPG varies depending on whether infection involves the

upper urinary tract or there is concomitant disease. Dipstick analysis often
(but not always) reveals hematuria and proteinuria. Leukocyte esterase
(WBCs) and nitrite (bacteria) test pads are not reliable in dogs and cats and
should not be used

Examination of urine sediment should be a routine part of a complete

urinalysis. Signiﬁcant numbers of WBCs (more than three to ﬁve per high-
power ﬁeld), often associated with hematuria or proteinuria, in a properly
collected urine sample suggest inﬂammation. Detection of signiﬁcant
microburia with pyuria indicates active inﬂammation associated with an
infection. Bacteria and fungi may be diﬃcult to identify in dilute urine,
making diagnosis of UTI problematic. In addition, UTI may be present
without concurrent inﬂammation if host defenses are compromised (eg,
hyperadrenocorticism, feline leukemia virus infection)

[7,11,25–29]

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

Rod-shaped bacteria may be identiﬁed in unstained preparations of urine

sediment if there are greater than 10,000 bacteria per milliliter of urine; these
organisms may not be consistently detected if present in fewer numbers.
Cocci are diﬃcult to detect in urine sediment if their numbers are less than
100,000 bacteria per milliliter

. Although detection of bacteria on urine

sediment examination suggests bacterial UTI, it should be veriﬁed by urine
culture. Urine sediment may be stained with Gram’s stain or new methylene
blue to aid in detection of microbes; however, urine culture is the ‘‘gold
standard’’ for conﬁrmation of UTI. Failure to detect bacteria on examina-
tion of urine sediment does not exclude their presence or rule out UTI.

Urine collection

For urine culture, urine should be collected by cystocentesis. With lower

urinary tract disease, collection of a sample by this method may be diﬃcult
because of pollakiuria; therefore, it may be necessary to collect urine for
culture by catheterization or, less desirably, during voiding. For these
techniques, the external genitalia of the patient should be cleaned.
Perivulvar fur may require clipping to prevent contamination. Although
urinary catheterization of dogs is usually accomplished without chemical
restraint, cats require sedation or anesthesia. Use a sterile catheter and
collection container (syringe or collection cup with a tight-ﬁtting lid). If
results of quantitative culture of urine samples obtained by catheterization
or midstream voiding are equivocal after serial cultures, collect urine by
cystocentesis.

The presence of bacteria in urine collected aseptically by cystocentesis,

even in low numbers, indicates UTI; however, false-positive results may
occur if a loop of intestine is penetrated with the hypodermic needle during
the procedure or if the sample is contaminated during handling. Contam-
ination usually involves recovery of more than one organism.

Urine culture

Quantitative urine culture is the gold standard for diagnosing UTI. A

diagnosis of UTI based only on clinical signs (eg, discolored urine,
pollakiuria, stranguria) or the presence of urinary tract inﬂammation is
a misdiagnosis that may result in inadequate or inappropriate treatment.
There are circumstances where antimicrobial therapy may be initiated
without results of urine culture; however, samples for urine culture should
be collected before initiating therapy. If antimicrobial therapy has been
started, it should be discontinued for 3 to 5 days before performing urine
culture to minimize inhibition of microbial growth.

Urine culture is the most deﬁnitive means of diagnosing a bacterial UTI.

Fungal culture should be considered when yeast or fungi are identiﬁed on
urine sediment examination. Care must be taken to collect, preserve, and

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

transport the urine sample appropriately to avoid contamination or
proliferation or death of bacteria

. Urine specimens for aerobic bacterial

culture should be transported and stored in sealed and sterilized containers,
and processing should begin as soon as possible. If laboratory processing is
delayed by more than 30 minutes, refrigerate the specimen (4(C)

. At

room temperature, bacterial counts may double every 20 to 45 minutes.
Multiplication or destruction of bacteria may occur within an hour of
collection.

If urine samples cannot be processed immediately for urine culture, there

are several alternatives available. Blood agar and MacConkey agar plates
may be inoculated and incubated for 24 hours. Use a calibrated bacterio-
logic loop or microliter mechanical pipette that delivers exactly 0.01 or 0.001
mL of urine to the culture plates. Streak the urine over the plates by
conventional methods. Blood agar supports growth of most aerobic
uropathogens, and MacConkey agar provides information that aids in
identiﬁcation of bacteria and prevents ‘‘swarming’’ of Proteus spp. Plates
can be stored in an incubator or under an incandescent light

. A 60-W

incandescent light positioned approximately 11 cm from the agar plate can
be used to maintain a surface temperature of approximately 35(C

. If

bacterial growth occurs on the plate after 24 hours, the plate may be
submitted for identiﬁcation and determination of antimicrobial susceptibil-
ity using the agar disk diﬀusion method

[32,33]

. If no growth occurs after 24

hours, plates may be discarded, because most organisms that cause UTI are
unlikely to grow after this time.

Commercially available urine culture collection tubes containing pre-

servative combined with refrigeration may be used to preserve specimens for
up to 72 hours

. More recently, in-house urine culture and susceptibility

kits have become available (IndicatoRx; IDEXX Laboratories, Westbrook,
ME).

Qualitative urine culture

A qualitative urine culture involves isolating and identifying bacteria in

urine; it does not include quantifying bacterial numbers. Although urine in
the bladder is normally sterile, urine that passes through the distal
urogenital tract often becomes contaminated with resident ﬂora (

Table 2

Therefore, interpretation of bacteria in urine collected by catheterization or
voiding is often diﬃcult to interpret even with quantiﬁcation of bacteria
(

Table 3

). For this reason, a diagnostic urine culture should include quanti-

tation of bacterial numbers in addition to identiﬁcation of the organism and
antimicrobial susceptibility.

Quantitative urine culture

A quantitative urine culture includes isolation and identiﬁcation of the

organism, and determination of the number of bacteria (colony-forming
units per unit volume). Quantitation of bacteria enables interpretation as to

929

J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

the signiﬁcance of bacteria present in a urine sample. Caution should be
exercised when interpreting quantitative urine cultures obtained by midstream
voiding or manual expression of urine. Although urine obtained from most
dogs without UTI was either sterile or contained less than 10,000 colony-
forming units per milliliter of urine (see

Table 3

), counts of 100,000 colony-

forming units or more per milliliter of urine occurred with suﬃcient frequency
to make collection of urine by these methods unsatisfactory

.The deﬁnition

of signiﬁcant bacteruria in cats involves lower numbers or organisms, because
cats seem to be more resistant to UTI than dogs and human beings.

Antimicrobial susceptibility testing

Administration of antimicrobial agents is the cornerstone of treating

UTI. The antimicrobial agent selected should be (1) easy to administer, (2)
associated with few if any side-eﬀects, (3) inexpensive, (4) able to attain
tissue or urine concentrations that exceed the minimum inhibitory concen-
tration (MIC) for the uropathogen by at least fourfold, and (5) unlikely to
aﬀect the patient’s intestinal ﬂora adversely

. Choice of antimicrobial

agent is based on antimicrobial susceptibility testing.

Table 2
Bacteria detected in the urogenital tract of normal male and female dogs

Male dogs

Female dogs

Genus

Distal urethra

Prepuce

Vagina

Acinetobacter

Bacteroides

Bacillus

Citrobacter

Corynebacterium

Enterococcus

Enterobacter

Escherichia

Flavobacterium

Haemophilus

Klebsiella

Micrococcus

Moraxella

Mycoplasma

Neisseria

Pasteurella

Proteus

Pseudomonas

Staphylococcus

Streptococcus

Ureaplasma

Data from

Barsanti JA, Johnson CA. Genitourinary infections. In: Greene CE, editor.

Infectious diseases of the dog and cat. 2nd edition. Philadelphia: WB Saunders; 1990. p. 157–83.

930

J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

Agar disk diﬀusion technique

Antimicrobial susceptibility testing often is done using agar disk diﬀusion

(Kirby-Bauer)

, which is adequate in most bacterial UTI. The agar disk

diﬀusion method uses Mueller-Hinton agar plates that have been inoculated
with a standardized suspension of a single uropathogen. Paper disks im-
pregnated with diﬀerent antimicrobial drugs are placed on the plate.
Eighteen to 24 hours after inoculation and incubation at 38(C, antimicro-
bial susceptibility is estimated by measuring zones of inhibition of bacterial
growth surrounding each disk. Zones of inhibition are then interpreted in
light of established standards, and susceptibility is recorded as resistant,
susceptible, or intermediate. Because of diﬀerences in the ability of various
antimicrobials to diﬀuse through agar, the antimicrobial disk surrounded by
the largest zone of inhibition is not necessarily the drug most likely to be
eﬀective. Also, because concentration of antimicrobial (except nitrofuran-
toin) in paper disks is comparable to typical serum concentration of the
drug, drugs that are found to be resistant by the agar disk diﬀusion method
may be eﬀective in the urinary tract if they are excreted in high concen-
trations in urine (eg, ampicillin, cephalexin).

Antimicrobial dilution technique

Antimicrobial dilution susceptibility tests are designed to determine the

minimum concentration of an antimicrobial drug that inhibits growth of the
uropathogen (ie, MIC). After inoculation and incubation of uropathogens
into wells containing serial twofold dilutions of antimicrobial drugs at
concentrations achievable in tissues and urine, MIC is deﬁned as the lowest
antimicrobial concentration (or highest dilution) that allows no visible
bacterial growth. The MIC is several dilutions lower than the minimum

Table 3
Interpretation of quantitative urine cultures in dogs and cats

(colony-forming units per

milliliter of urine)

Signiﬁcant

Suspicious

Contaminant

Dogs

Cats

Dogs

Cats

Dogs

Cats

Cystocentesis

1000

100–1000

100

Catheterization

10,000

1000

1000–10,000

100–1000

1000

100

Midstream voiding

100,000

10,000 10,000–90,000 1000–10,000 10,000 1000

Manual compression

100,000

10,000 10,000–90,000 1000–10,000 10,000 1000

These data represent generalities. Occasionally, bacterial urinary tract infection may exist

with fewer organisms (ie, false-negative results).

Contamination of midstream voided samples may cause growth of

10,000 colony-

forming units per milliliter (ie, false-positive results); therefore, these samples should not be used
routinely for diagnostic culture.

Data from

Lulich JP, Osborne CA. Bacterial urinary tract infections. In: Ettinger SJ,

Feldman EC, editors. Textbook of veterinary internal medicine. 4th edition. Philadelphia: WB
Saunders; 1999. p. 1775–1788.

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

bactericidal concentration of drugs. In general, antimicrobial agents are
likely to be eﬀective if they can achieve a urine concentration that is 4 times
the MIC. Many antimicrobial drugs that are renally excreted reach concen-
trations in urine that are 10 to 100 times greater than serum concentrations.

Summary

UTI is an important cause and consequence of urinary tract disease and

may occur in association with systemic disease. Diagnosis of UTI is not
diﬃcult and is essential for providing optimal patient care. Furthermore,
appropriate diagnostic testing is needed to avoid inappropriate treatment of
patients (eg, undertreatment, overtreatment).

References

[1] Ling GV. Therapeutic strategies involving antimicrobial treatment of the canine urinary

tract. J Am Vet Med Assoc 1984;185(10):1162–4.

[2] Lulich JP, Osborne CA. Bacterial urinary tract infections. In: Ettinger SJ, Feldman EC,

editors. Textbook of veterinary internal medicine. 4th edition. Philadelphia: WB Saunders;
1999. p. 1775–88.

[3] Lulich JP, Osborne CA, Bartges JW, Lekcharoensuk C. Canine lower urinary tract

disorders. In: Ettinger SJ, Feldman EC, editors. Textbook of veterinary internal medicine,
vol. 2. 5th edition. Philadelphia: WB Saunders; 1999. p. 1747–83.

[4] Lees GE. Bacterial urinary tract infections. Vet Clin North Am Small Anim Pract 1996;

26(2):297–304.

[5] Thomsen MK, Svane LC, Poulsen PH. Canine urinary tract infection. Detection, pre-

valence and therapeutic consequences of bacteriuria. Nord Vet Med 1986;38(6):394–402.

[6] Lees GE. Epidemiology of naturally occurring feline bacterial urinary tract infections. Vet

Clin North Am Small Anim Pract 1984;14(3):471–9.

[7] Bartges JW, Blanco L. Bacterial urinary tract infection in cats. Compend Standard Care

2001;3:1–5, 9.

[8] Lekcharoensuk C, Osborne CA, Lulich JP. Epidemiologic study of risk factors for lower

urinary tract diseases in cats. J Am Vet Med Assoc 2001;218(9):1429–35.

[9] Lulich JP, Osborne CA. Fungal infections of the feline lower urinary tract. Vet Clin North

Am Small Anim Pract 1996;26(2):309–15.

[10] Ling GV, Norris CR, Franti CE, et al. Interrelations of organism prevalence, specimen

collection method, and host age, sex, and breed among 8,354 canine urinary tract
infections (1969–1995). J Vet Intern Med 2001;15(4):341–7.

[11] Pressler BM, Vaden SL, Lane IF, Cowgill LD, Dye JA. Candida spp. urinary tract

infections in 13 dogs and seven cats: predisposing factors, treatment, and outcome. J Am
Anim Hosp Assoc 2003;39(3):263–70.

[12] Fulton R, Jr, Walker RD. Candida albicans urocystitis in a cat. J Am Vet Med Assoc 1992;

200(4):524–6.

[13] Kruger JM, Osborne CA. The role of viruses in feline lower urinary tract disease. J Vet

Intern Med 1990;4(2):71–8.

[14] Kruger JM, Osborne CA, Goyal SM, Pomeroy KA, O’Brien TD. Clinicopathologic and

pathologic ﬁndings of herpesvirus-induced urinary tract infection in conventionally reared
cats. Am J Vet Res 1990;51(10):1649–55.

[15] Kruger JM, Osborne CA, Goyal SM, O’Brien TD, Pomeroy KA, Semlak RA.

Clinicopathologic analysis of herpesvirus-induced urinary tract infection in speciﬁc-
pathogen-free cats given methylprednisolone. Am J Vet Res 1990;51(6):878–85.

932

J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

[16] Kruger JM, Osborne CA. The role of viruses in feline lower urinary tract disease. J Vet

Intern Med 1990;4(2):71–8.

[17] Kruger JM, Venta PJ, Swenson CL, et al. Prevalence of bovine herpesvirus-4 infection in

cats in Central Michigan. J Vet Intern Med 2000;14(6):593–7.

[18] Crawford JT, Adams WM. Inﬂuence of vestibulovaginal stenosis, pelvic bladder, and

recessed vulva on response to treatment for clinical signs of lower urinary tract disease in
dogs: 38 cases (1990–1999). J Am Vet Med Assoc 2002;221(7):995–9.

[19] Osborne CA, Caywood DD, Johnston GR, et al. Feline perineal urethrostomy: a potential

cause of feline lower urinary tract disease. Vet Clin North Am Small Anim Pract 1996;
26(3):535–49.

[20] Adams WM, DiBartola SP. Radiographic and clinical features of pelvic bladder in the dog.

J Am Vet Med Assoc 1983;182(11):1212–7.

[21] Lavocat MP, Granjon D, Allard D, Gay C, Freycon MT, Dubois F. Imaging of

pyelonephritis. Pediatr Radiol 1997;27(2):159–65.

[22] Rushton HG, Majd M. Dimercaptosuccinic acid renal scintigraphy for the evaluation of

pyelonephritis and scarring: a review of experimental and clinical studies. J Urol 1992;
148(5 Part 2):1726–32.

[23] Buﬃngton CA, Chew DJ, Kendall MS, DiBartola SP. Clinical evaluation of cats with

nonobstructive urinary tract disease. J Am Vet Med Assoc 1997;210:46–50.

[24] Vail DM, Allen TA, Weiser G. Applicability of leukocyte esterase test strip in detection of

canine pyuria. J Am Vet Med Assoc 1986;189(11):1451–3.

[25] Gerding PA, Jr, Morton LD, Dye JA. Ocular and disseminated candidiasis in an

immunosuppressed cat. J Am Vet Med Assoc 1994;204(10):1635–8.

[26] Forrester SD, Troy GC, Dalton MN, Huﬀman JW, Holtzman G. Retrospective evaluation

of urinary tract infection in 42 dogs with hyperadrenocorticism or diabetes mellitus or
both. J Vet Intern Med 1999;13(6):557–60.

[27] Barsanti JA, Brown J, Marks A, Reece L, Greene CE, Finco DR. Relationship of lower

urinary tract signs to seropositivity for feline immunodeﬁciency virus in cats. J Vet Intern
Med 1996;10(1):34–8.

[28] Kruger JM, Osborne CA, Goyal SM, et al. Clinical evaluation of cats with lower urinary

tract disease. J Am Vet Med Assoc 1991;199(2):211–6.

[29] Buﬃngton CA, Chew DJ, Kendall MS, et al. Clinical evaluation of cats with

nonobstructive urinary tract diseases. J Am Vet Med Assoc 1997;210(1):46–50.

[30] Padilla J, Osborne CA, Ward GE. Eﬀects of storage time and temperature on quantitative

culture of canine urine. J Am Vet Med Assoc 1981;178(10):1077–81.

[31] Carter JM, Klausner JS, Osborne CA, Bates FY. Comparison of collection techniques for

quantitative urine culture in dogs. J Am Vet Med Assoc 1978;173(3):296–8.

[32] Saunders A, Bartges JW, Bemis DA, Bryant MJ, Duckett R. Evaluation of blood agar

plates and incandescent lighting for aerobic bacterial urine cultures [abstract]. J Vet Intern
Med 2002;16:379.

[33] Blanco LJ, Bartges JW, New J, Bemis DA, Bryant MJ, Duckett R. Evaluation of blood

agar plates as a transport medium for aerobic bacterial urine cultures [abstract]. J Vet
Intern Med 2001;15:303.

[34] Allen TA, Jones RL, Purvance J. Microbiologic evaluation of canine urine: direct

microscopic examination and preservation of specimen quality for culture. J Am Vet Med
Assoc 1987;190(10):1289–91.

[35] Barsanti JA, Johnson CA. Genitourinary infections. In: Greene CE, editor. Infectious

diseases of the dog and cat. 2nd edition. Philadelphia: WB Saunders; 1990. p. 157–83.

[36] Osborne CA, Lees GE. Bacterial infections of the canine and feline urinary tract. In:

Osborne CA, Finco DR, editors. Canine and feline nephrology and urology. Baltimore:
Williams & Wilkins; 1995. p. 759–97.

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J.W. Bartges / Vet Clin Small Anim 34 (2004) 923–933

Veterinary hemodialysis: advances in

management and technology

Julie R. Fischer, DVM

*, Valeria Pantaleo, Dr Vet Met

Thierry Francey, Dr Vet Med

c,d

Larry D. Cowgill, DVM, PhD

a,d

University of California Veterinary Medical Center at San Diego, PO Box 9415,

6525 Calle del Nido, Rancho Santa Fe, CA, USA

Veterinary Medical Teaching Hospital, School of Veterinary Medicine,

University of California at Davis, Davis, CA, USA

Companion Animal Hemodialysis Unit, School of Veterinary Medicine,

University of California at Davis, Davis, CA, USA

Department of Medicine and Epidemiology, School of Veterinary Medicine,

University of California at Davis, Davis, CA, USA

Hemodialysis (HD) is a renal replacement therapy that provides a bridge

of metabolic stability to patients who would otherwise die from the
pansystemic ramiﬁcations of severe uremia. This therapeutic option has
been consistently available on a geographically limited basis to the
veterinary community since the ﬁrst service opened in 1990 at the University
of California at Davis, Veterinary Medical Teaching Hospital. Since that
time veterinary experience with this treatment modality has deepened and
reﬁned as the caseload and number of veterinarians trained in HD have
increased and as relevant technologic advances have been incorporated into
the practice of veterinary dialysis. Currently, ﬁve facilities in North America
oﬀer HD for companion animals on both an emergent and chronic basis
(

Appendix A

), and the program at the University of California at Davis

provides postresidency training in nephrology and renal replacement
modalities. As awareness of the usefulness and availability of dialytic
therapy increases among veterinarians and pet owners and the number of
veterinary dialysis facilities increases, dialytic management will become the
standard of advanced care for animals with severe intractable uremia.

* Corresponding author.
E-mail address:

jroﬁscher@ucdavis.edu

(J.R. Fischer).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.007

Vet Clin Small Anim

34 (2004) 935–967

Principles, applications, and advances in small animal hemodialysis

Principles of hemodialysis

Strictly speaking, dialysis refers to the net movement of solutes and water

across a semipermeable membrane along concentration gradients. HD is the
extracorporeal exchange of water and solutes between blood and a contrived
solution termed dialysate across manufactured semipermeable membranes
for the purpose of removing metabolic waste products and correcting the
ﬂuid, electrolyte, and acid-base derangements that renal failure eﬀects
(

). The dialysate is formulated to favor movement of permeable waste

molecules (eg, urea, creatinine) out of the plasma, to maintain physiologic
plasma concentrations of permeable substances (eg, glucose, phosphorus,
calcium), and to replenish or load permeable molecules that have been
depleted from the plasma (eg, bicarbonate). The three physical principles
governing solute and ﬂuid waste removal during HD are diﬀusion, ultraﬁl-
tration, and convection

Diﬀusive dialysis depends on the random molecular motion of dissolved

particles. As these particles arbitrarily encounter pores in the dialyzer
membrane, they move from one side of the membrane to the other through
membrane channels (

). Likelihood of contact with a membrane channel

is directly proportional to the concentration of a given particle type and its
thermodynamic energy. Thermodynamic energy is inversely proportional to

SCHEMATIC CROSS-

SECTION OF A

DIALYZER FIBER

DIALYSATE

DIALYSIS
MEMBRANE

BLOOD

DIALYSIS DELIVERY

DIALYZER

(ARTIFICIAL KIDNEY)

SYSTEM

Fig. 1. (Left) A Cobe Centrysystem C3 Plus dialysis delivery system with a dialyzer in place.
(Middle) An enlarged view of a hollow-ﬁber dialyzer; the dialysate inﬂow and outﬂow ports are
on the left, and the blood inﬂow and outﬂow ports are on the right. (Right) A schematic cross
section of a single dialyzer ﬁber. Blood ﬂows through the longitudinally oriented hollow ﬁbers,
and the dialysate circulates around them within the plastic casing.

936

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

molecular mass; thus, at equal concentrations, smaller molecules diﬀuse more
readily than larger molecules. If concentrations of a solute become equal on
both sides of the membrane (ie, ﬁltration equilibrium is achieved), diﬀusion
across the membrane still occurs but net transfer of that solute is zero.
Maintenance of the concentration gradient between blood and dialysate, and
therefore maintenance of diﬀusive dialysis, is accomplished by continuous
replacement of dialysate, thus preventing ﬁltration equilibrium

The peristaltic blood pump on the dialysis machine acts as an

extracorporeal ‘‘heart,’’ forcing the patient’s blood through external tubing
to the dialyzer and generating an outwardly directed hydrostatic force
across the dialyzer membrane. This hydrostatic force provides the impetus
for ﬂuid transfer across the membrane by ultraﬁltration, analogous to the
ultraﬁltration that occurs in the glomerulus. Counterpressure on the
dialysate side of the membranes prevents hydraulic transfer of ﬂuid from
the plasma to the dialysate. To facilitate and regulate ultraﬁltration, the
outward transmembrane pressure generated by the blood pump is compli-
mented by application of a vacuum to the dialysate side of the membrane.
An outward net hydraulic pressure draws water molecules from the blood
through the dialyzer membrane pores and into the dialysate (

). The

amount of water that can be moved across the membrane during a given
time depends on the hydraulic permeability of the dialysis membrane, the
membrane surface area, and the hydrostatic gradient across the membrane.

SEMI-PERMEABLE

MEMBRANE

BLOOD

HCO

DIALYSATE

HCO

bicarbonate

urea

Urea
Creatinine

creatinine

Fig. 2. Schematic representation of diﬀusion between blood and dialysate across a dialyzer
membrane. Solutes diﬀuse through the dialyzer membrane pores across the membrane in both
directions; the arrows represent the direction of net diﬀusion of solutes according to
concentration gradients.

937

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

This value is represented by the ultraﬁltration coeﬃcient (KUf), which
quantiﬁes milliliters of ﬂuid that can be transferred per millimeter of
mercury transmembrane pressure per hour

During ultraﬁltration, solute particles dissolved in plasma water are

pulled through the dialyzer membrane pores along with the water molecules.
This ‘‘solvent drag,’’ or convection, contributes signiﬁcantly to the total
solute load removed during a dialysis session. Convective solute removal
occurs independent of diﬀusive gradients and particularly enhances the
removal of middle-molecular-weight substances (500–15,000 d) that are less
eﬃciently removed by diﬀusive dialysis. Hemoﬁltration, a process per-
formed alone or as part of a dialysis treatment, exploits this convective
principle and consists of infusing large volumes of isotonic intravenous
replacement ﬂuid while simultaneously removing an equal volume of plasma
water via ultraﬁltration. When performed concomitant with HD, the
process is termed hemodiaﬁltration and maximizes the eﬀects of diﬀusive
as well as convective dialysis

[1–4]

Veterinary applications for hemodialysis

As outlined in

, HD provides therapeutic beneﬁt to three broad

categories of veterinary patients: (1) animals with severe uremia and its

DIALYSATE

SEMI-PERMEABLE

MEMBRANE

VACUUM

PRESSURE

water

solutes

Urea
Creatinine

BLOOD

Fig. 3. Schematic representation of ultraﬁltration and convection between blood and dialysate
across a dialyzer membrane. Positive pressure on the blood side and negative pressure on the
dialysate side of the dialysis membrane combine to draw water and solute molecules through
the membrane pores in processes called ultraﬁltration and convection, respectively. Note that
the concentration of dissolved solute in the blood remains the same in these processes, because
solutes and water are simultaneously moved through the pores. The arrow represents the
direction of net movement of solute and water.

938

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

component metabolic derangements, (2) animals with intractable volume
overload, and (3) animals with some toxicoses. Most animals presented for
HD are acutely uremic and nonresponsive to attempted diuresis with
intravenous ﬂuids and pharmacologic manipulation. The uremia arises from
an acute renal insult or an acute exacerbation of an underlying renal disease.
HD can also mitigate the clinical manifestations of chronic end-stage renal
disease when conventional management fails, but few owners are ﬁnancially
able to continue HD indeﬁnitely. Life-threatening volume overload, whether
caused by oliguria/anuria, congestive heart failure, or excessive ﬂuid
administration, can be managed with ultraﬁltration in a patient that is

Box 1. Applications for hemodialysis therapy in dogs and cats

Severe uremia

Acute renal failure

Refractory azotemia (blood urea nitrogen [BUN]

100 mg/dL

or creatinine

10 mg/dL)

Severe electrolyte disturbance (hyperkalemia, hypo/

hypernatremia)

Severe metabolic acidosis
Management of delayed graft function after transplantation

Chronic (end-stage) renal failure

Refractory azotemia (BUN

100 mg/dL or creatinine 10

mg/dL)

Preoperative conditioning for renal transplantation
Finite extension of improved quality of life to allow client

adjustment to diagnosis and prognosis

Volume overload

Unresponsive oligoanuria
Fulminant congestive heart failure

Pulmonary edema
Circulatory overload
Lack of response to diuretics

Iatrogenic fluid overload
Parenteral nutrition in oligoanuric animals

Acute toxicosis or drug overdose

Ethylene glycol toxicosis (acute toxin removal and chronic

management of resultant ARF)

Environmental/agricultural toxins
Accidental ingestion/overdose of many medications (aspirin,

acetaminophen, phenylbutazone, digoxin, amikacin,
azathioprine, cyclophosphamide, enalapril, procainamide,
phenobarbital, and theophylline among others)

939

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

nonresponsive to diuretics. Finally, dialytic techniques are uniquely suited
to the management of speciﬁc acute toxicoses. As outlined in

, drugs

and chemicals whose physical characteristics permit passage through
dialyzer membrane pores and which are not bound to plasma proteins
can be quickly and eﬃciently removed from the bloodstream, often with
a single HD session. Referral guidelines for HD patients are presented in

Appendix B

Advances in vascular access

Establishment of reliable and durable high-volume vascular access is an

essential requirement for HD and a frequent cause of frustration and
therapeutic compromise for human and veterinary dialysis clinicians alike

[2,5,6]

. The two currently available methods of HD vascular access in small

animals are transcutaneous venous catheters and surgically created arterio-
venous ﬁstulas.

Catheters

Large-gauge intravenous catheters are the current standard for vascular

access in animals undergoing HD

[2,7,8]

. Dialysis catheters must allow

suﬃcient blood ﬂow to permit delivery of blood to the dialyzer at a rate of 5
to 10 mL/kg/min or greater. For example, in a cat weighing 4 kg, the dialysis
catheter should permit blood treatment at a rate of at least 1.2 L/h or 20
mL/min. In patients weighing less than 10 kg, 7-French dual-lumen
catheters, such as those used for blood sampling and parenteral nutrition,
may be used on a short-term basis. Catheters need to be at least 10 cm long
for even the smallest dialysis patients; however, most animals require
a catheter 15 cm in length or greater. For patients weighing more than 10 kg,
11.5-French, silicone-based, temporary human dialysis catheters (Hemocath
11.5-French, 24-cm, dual-lumen catheter; Medcomp, Harleysville, PA) are
used commonly for acute access (

). Placement of catheters in emergent

situations can usually be accomplished with local anesthesia and light
tranquilization via a percutaneous modiﬁed Seldinger (over-the-wire)
technique

provided that the jugular vein is not severely traumatized. In

patients with signiﬁcant prior jugular trauma from venipuncture or
catheterization, expedient catheter placement may require a surgical ap-
proach and venotomy.

Acute catheters generally provide adequate blood ﬂow for 2 to 4 weeks

[2,5]

. In patients whose dialytic therapy extends longer than the acute

catheter’s functional life, a permanent subcutaneously tunneled dialysis
catheter is usually placed (

). Many diﬀerent types and sizes of

permanent catheters are marketed for HD; most are silicone based or
modiﬁed and softened polyurethane. Most catheters are dual lumen in
design (eg, Permcath; Quinton Instruments Co., Seattle, WA), but some
consist of two single-lumen catheters placed through separate venotomy

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Box 2. Readily dialyzable medications and chemicals

Alcohols

Ethanol
Ethylene glycol
Methanol

Analgesics/anti-inflammatories

Acetaminophen
Aspirin
Mesalamine (5-ASA)
Morphine

Pentazocine

Antibacterials

Amikacin
Amoxicillin (most penicillins)
Cephalexin (most first-generation cephalosporins)
Cefotetan (many second-generation cephalosporins)
Cefoxitin
Ceftriaxone (many third-generation cephalosporins)
Chloramphenicol
Gentamicin
Imipenem/cilastatin
Kanamycin
Linezolid
Nitrofurantoin
Ofloxacin
Metronidazole
Sulbactam
Sulfamethoxazole
Sulfisoxazole
Trimethoprim
Vancomycin

Anticonvulsants

Gabapentin
Phenobarbital
Phenytoin

Primidone

Antifungals

Dapsone
Fluconazole
Flucytosine

Antineoplastics

Busulfan
Carboplatin

(continued on next page)

941

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Cytarabine

Cyclophosphamide
Fluorouracil (5-FU)
Ifosfamide
Methotrexate
Mercaptopurine

Antivirals

Acyclovir
Famciclovir
Valacyclovir
Zidovudine

Cardiac/vasoactive medications

Atenolol
Bretylium
Captopril
Enalapril
Esmolol
Lisinopril
Metoprolol
Mexiletine
Nitroprusside
Procainamide
Sotalol
Tocainide

Chelating agents

Deferoxamine
Ethylenediamine tetraacetic acid (EDTA)
Penicillamine

Immunosuppressive agents

Azathioprine
Methyl prednisone

Miscellaneous medications

Allopurinol
Ascorbic acid
Carisoprodol
Chloral hydrate
Chlorpheniramine
Diazoxide
Foscarnet
Iohexol
Iopamidol
Lithium
Mannitol
Metformin

942

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

sites in the same vessel and have two separate subcutaneous tunnels (Tesio
Cath, Medcomp). A recent design advance is the development of split-tip
catheters, the ﬁrst of which was the Ash Split Cath (Medcomp). Split-tip
catheters are dual lumen for much of their length, but separate into two
single-lumen catheters at the distal aspect. This separation allows the
individual sides to move freely within the vessel, helping to prevent
occlusion of the inﬂow portal as a result of aspiration against the vessel
wall and decreasing recirculation when compared with ﬁxed-tip catheters

. The split-tip design permits the high ﬂows achieved by the Tesio Cath

but requires only a single tunnel and venotomy, facilitating placement

Common to all permanent catheters is the presence of a small Dacron cuﬀ
encircling the exterior of the catheter. When properly placed, this cuﬀ lies in
the subcutaneous tunnel between the venotomy site and the site of skin
penetration. Fibroblasts migrate into the cuﬀ matrix, forming a connective
tissue bond between the subcutis and the catheter. This tissue bond anchors
the catheter, stabilizing the placement of the catheter tip and diminishing the
chance of dislodgement from the vessel. It also provides a physical barrier
between the skin exit site and the venotomy site, helping to prevent local
extension of tunnel infections into the vasculature

[10,11]

Fistulas

Surgical arteriovenous ﬁstulas and arteriovenous polytetraﬂuoroethylene

grafts are the mainstays of vascular access in human HD patients and are
ideally created subcutaneously in the forearm

. Fistulas demonstrate

greater durability and fewer complications than other methods of angioac-
cess and are the National Kidney Foundation’s Kidney Disease Outcomes
Quality Initiative’s recommended ﬁrst-choice access

[11–16]

. To create

a ﬁstula, an artery and vein (preferably the radial artery and cephalic vein)
are surgically anastomosed; ﬁstula maturation usually occurs over 6 to 8
weeks but can take several months. The increased pressure and ﬂow from
anastomosis with the artery cause dilation of the vein, creating a readily
visible and palpable vascular segment that is punctured percutaneously for
each HD treatment. Large-bore (14-gauge) needles are inserted in the vessel
toward the arterial and venous blood ﬂows and are connected to the arterial

Minoxidil
Octreotide
Ranitidine
Theophylline

High-flux dialysis only.

If hemodialysis is instituted while blood concentrations are still high, these

substances can be substantially cleared.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

and venous blood lines for withdrawal and return of blood, respectively.
Fistulas have not been used routinely for companion animal HD, but
a recent study by Adin et al

investigated the suitability of three

anatomic sites for creation of ﬁstulas for use in HD procedures in dogs. In
Adin et al’s study

, brachial artery–cephalic vein ﬁstulas matured well

and permitted adequate access, needle stabilization, and blood ﬂow after
a 56-day maturation period

[15,17]

. Given the longer performance and lower

incidence of thrombosis and infection associated with ﬁstulas compared
with catheters in human dialysis patients, this access method merits further
clinical investigation in canine HD patients undergoing long-term therapy.

Subcutaneous vascular ports

A fully implantable, titanium alloy, subcutaneous vascular access device

(LifeSite Hemodialysis Access System; Vasca, Tewksbury, MA) has been in
use in human patients for several years; in one study, increased blood ﬂow
and decreased incidence of thrombosis and catheter-related infection were
demonstrated compared with tunneled and cuﬀed dialysis catheters

[18,19]

Fig. 4. Temporary hemodialysis (HD) catheters. (A) A 7-French, 20-cm, dual-lumen catheter
commonly used for parenteral nutrition and blood sampling. (B) An 11.5-French, 24-cm, dual-
lumen catheter designed for temporary HD access in human beings. (C) A peripheral catheter
(used for gaining initial venous access), vein dilator, and guidewire coiled in its holder (top to
bottom

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

The LifeSite system is similar to the subcutaneous ports currently used in
veterinary medicine for chronic chemotherapy and repeated blood collec-
tions but has titanium alloy valves instead of the silicone rubber septum of
the veterinary device. The metal valves permit more secure locking of the
access needle in the device than is possible with the silicone septum;
additionally, because the valve parts to admit the needle rather than being
punctured, the valve does not fatigue as rapidly as the silicone septum does.
For HD, two separate ports are implanted and a silastic cannula tunnels
subcutaneously from each access port to a venotomy site in the selected vein.
The LifeSite functions similarly to the twin-lumen transcutaneous catheters,
with independent conduits for blood outﬂow and return. The LifeSite
system is currently cost-prohibitive for veterinary medicine, but use of other
subcutaneous port systems, such as the Companion Port (Norfolk Veter-
inary Products, Skokie, IL), merits investigation. Implantation of such ports
for HD use would expand potential access sites (eg, femoral veins, brachial
veins, caudal vena cava) and could decrease the incidence of bacterial
infection and thrombosis seen with transcutaneous catheters.

Fig. 5. Three types of permanent hemodialysis catheter. (A) Pediatric (middle) and adult (right)
dual-lumen ﬁxed-tip permanent catheters. On the far left is a peel-away sheath introducer for
percutaneous placement. (B) Tesio Cath twin catheters. An access port has been connected to
only one of the catheters in this picture. Below the catheters are tunneling devices and peel-away
sheaths. (C) Ash Split catheter. This catheter combines the ease of ﬁxed-tip placement with the
enhanced ﬂows of twin catheters. A tunneling device, vein dilator, and peel-away sheath are
pictured to the right of the catheter. Note the Dacron cuﬀ at the proximal end of all permanent
catheters.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Locking solutions

During the interdialytic interval, dialysis catheters are routinely locked

with varying concentrations (100–5000 U/mL) of heparin to prevent
intraluminal thrombosis. Concentration of the locking solution is de-
termined empirically based on the animal’s body weight and the anticipated
interval between dialysis treatments. Because small volumes of heparin leach
out of the catheter into the bloodstream, catheters in smaller patients are
locked with lower concentrations of heparin to avoid systemic overhepari-
nization. Catheter locks are changed at intervals of no greater than 1 week
to maintain patency.

Trisodium citrate is used primarily as an anticoagulant, but recent studies

investigating trisodium citrate as a catheter lock solution demonstrate
antimicrobial properties as well

[20–22]

. The Ca

and Mg

chelating

properties of trisodium citrate may account for its antimicrobial eﬀect by
preventing luminal bioﬁlm formation

[20,21]

. Bioﬁlms consist of microbial

communities proliferating on surfaces in an aqueous environment and
generally form within 1 to 14 days of catheter insertion

. These

communities elaborate a coating of exopolysaccharide, which enables ﬁrm
surface adhesion for bacterial progeny, and a glycocalyx matrix, which
envelops the colonies as they grow

. Persistence of the bioﬁlm renders

complete bacterial eradication extremely diﬃcult, establishing an infective
nidus that permits intermittent bacteremia and development of distant
infections.

Recent studies have also investigated the use of taurolidine, a biocom-

patible antimicrobial with potent broad-spectrum bactericidal properties
and some anticoagulant properties, in combination with citrate as a novel
catheter lock solution

[24,25]

. This combination was proposed to help

eradicate luminal bioﬁlms and thus to provide prophylaxis against
bacteremia while still preventing catheter clotting. Taurolidine shows
excellent eﬃcacy in reducing catheter-related bacteremia in human HD
patients

and seems to have the capacity to destroy preexisting luminal

bioﬁlms as well as to prevent their formation

. Allon

recently

demonstrated that the taurolidine-citrate lock solution was also associated
with a higher rate of catheter thrombosis, although Stas et al

found no

diﬀerence in luminal thrombus formation between 30% trisodium citrate
and heparin, 5000 U/mL. Because bacterial catheter infection is a pervasive
and modiﬁable agent of dialysis-related morbidity and mortality, trisodium
citrate and taurolidine as single agents and in combination warrant
evaluation for eﬃcacy and utility in veterinary patients.

Dialysis delivery systems

The core functions of the dialysis delivery system are (1) to generate

a prescribed dialysate and continuously monitor its composition, tempera-
ture, pH, and ﬂow; (2) to regulate and monitor the ﬂow of blood in the
extracorporeal circuit; (3) to regulate the volume and rate of ultraﬁltration;

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

(4) to deliver anticoagulant; and (5) to display the status of vital systems and
protect the patient from unsafe conditions

[7,26]

. Technologic advances in the

past 5 years have enhanced these core functions to improve the delivery and
safety of dialytic therapy to animals. Modern dialysis delivery systems have
incorporated sophisticated computer interfaces that help to eliminate the
mystique and complexity of HD prescriptions, display real-time trends of the
patient’s status and treatment progress, and incorporate biofeedback loops to
alter HD conditions interactively in response to changes in the physiology of
the patient. Touch screen technology and organization of dialysis functions
into display pages have simpliﬁed user interaction with the dialysis treatment
and facilitated ongoing evaluation of the patient’s clinical status.

Dialyzers and dialysate

Hollow-ﬁber dialyzers with ﬁbers constructed chieﬂy of cellulosynthetic

(eg, Hemophan, Gambro Renal Products, Lakewood, Colorado; Cellosyn)
or synthetic noncellulosic materials (eg, polyacrylonitrile, polysulfone,
polyamide, polymethylmethacrylate) are the current standard for veterinary
HD. Use of these membranes has increased biocompatibility and decreases
complement activation compared with earlier cellulose and substituted
cellulose membranes

[3,27]

. The development of cellulosynthetic and

synthetic membranes led to the advent of ‘‘high-ﬂux’’ dialyzers that have
larger pores, permitting better middle molecule removal and higher rates of
ultraﬁltration

[2,3]

. Initially cost-prohibitive for veterinary use, the price of

synthetic membrane dialyzers is now comparable to that of cellulosic devices,
warranting their routine use in animal patients.

The widespread use of highly permeable dialyzer membranes substantially

increases the exposure of dialysis patients to pyrogens, endotoxins, and
bacterial contaminants residing in the dialysate water source and delivery
piping These contaminants promote cytokine induction and inﬂammatory
responses that contribute to the morbidity associated with HD treatments

. New dialysis delivery systems provide a highly restrictive ultraﬁltration

step that generates ‘‘ultrapure’’ dialysate immediately before its delivery to
the dialyzer, thus decreasing endotoxin and bacterial contamination of
dialysate

. Additionally, many dialysis machine manufacturers now

provide systems that generate bicarbonate-based dialysate from powered
concentrates, eliminating the potential for the bacterial contamination that
has been seen with the use of liquid bicarbonate sources. Some newer delivery
systems also have integrated programs for routine chemical or heat
disinfection of the internal ﬂuid path within the machine to retard the
development of bioﬁlms and prevent inadvertent bacterial proliferation.

Monitoring modalities

Technologic advancements in the monitoring of treatment eﬀectiveness

and patient status have contributed greatly to global improvement in
dialysis delivery to uremic animals. Many external monitoring devices are

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

now integrated into the designs of new-generation delivery systems and thus
are available at every dialysis session. Classic blood-side assessments of
dialyzer and urea removal kinetics are conventional standards for quanti-
tating treatment eﬃcacy, but these parameters are conceptually diﬃcult and
problematic to measure and thus are rarely documented in animal patients.
To facilitate more accurate computation of dialyzer clearance and urea
kinetics, new delivery systems display estimates of compensated blood ﬂow
that account for the inﬂuence of negative arterial pressure on pump segment
eﬃciency and provide more accurate estimates of extracorporeal blood ﬂow
and thus of the blood volume processed.

Recent validation of ionic dialysance as a substitute for dialyzer urea

clearance (K

(urea)

) permits bloodless on-line kinetic modeling of dialysis

sessions on appropriately equipped machines. Dialysance is a measure of
solute mass transfer from blood to dialysate when the solute is present in the
blood and dialysate. The clearance of a solute across the dialyzer is equal to
its dialysance when the solute is present only in the blood and absent in the
dialysate. Ionic dialysance is a measure of the transfer characteristics of all
small-molecular-weight ions that contribute to the conductivity of the
patient’s plasma. Because the collective dialysance of these small-molecular-
weight ions is deemed generally similar to the dialysance of urea, ionic
dialysance serves as a reasonable surrogate for urea dialysance. In
a conventional single-pass HD circuit, urea dialysance becomes equal to
K

(urea)

; thus, ionic dialysance becomes an acceptable predictor of K

(urea)

and

the delivered dialysis dose (K

(urea)

t), where t is treatment time. Ionic

dialysance can be measured noninvasively, sequentially, and in real time
(without blood sampling) by measurement of the conductivity of the
dialysate at the inlet and outlet ports of the dialyzer before and in response
to programmed spikes in dialysate conductivity

[28–30]

. With regular

assessment during the dialysis session, ionic dialysance provides an in-
dividualized quality assurance measure of the delivered dialysis dose for
each treatment session. Furthermore, decreases in ionic dialysance during
the treatment predict real-time alterations in treatment eﬃcacy (eg, as
a result of dialyzer clotting, increased access recirculation, blood ﬂow
discrepancies) and thus provide an opportunity to intervene during the
treatment to ensure adequate therapy is administered. Because access
recirculation is incorporated in the kinetic algorithm deﬁning the ionic
mass transfer and ionic dialysance, these same methods can be used to
estimate access blood ﬂow rate and the recirculation ratio of the access when
the ionic dialysance measurements are made with the blood lines in the
normal and reversed positions. The availability and utility of ionic
dialysance in newer machines and the relative ease with which it predicts
dialysis delivery should promote a better understanding of the kinetics of
dialytic therapies and increase the eﬃcacy of dialysis prescriptions.

External in-line hematocrit monitors (Crit-Line III; Hema Metrics,

Kaysville, UT) have been available for blood volume monitoring and are

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

vital for hemodynamic assessment of animal patients during the HD
treatment

[7,26]

. This technology is now incorporated into the functions

and displays of some dialysis delivery systems. These systems provide real-
time quantitation and graphic display of the eﬀects of ultraﬁltration on blood
volume and help to predict adverse hemodynamic events. When integrated
into the structure of the delivery system, this monitoring function can be
combined with biofeedback systems to modify preset ultraﬁltration (or
sodium or ultraﬁltration proﬁling) prescriptions to limits that are safe and
tolerable. These integrated systems can alarm the system or stop ultraﬁltra-
tion when critical changes in blood volume occur. An additional biofeedback
system uses blood temperature sensors that interact with dialysate temper-
ature to detect and correct blood temperature increases. This feedback
system decreases the return blood temperature by changing dialysate
temperature to forestall progression or exacerbation of hypotensive or
vasoconstrictive events that can occur with blood temperature changes.

Blood pressure monitoring is also integrated into the hardware and

interactive display in new dialysis delivery systems. Although this feature
permits real-time display of the patient’s hemodynamic trends, the employed
oscillometric systems may not be as reliable for small dogs and cats as
devices designed for animals. In all, there have been major advances in the
design and functionality of HD equipment to ensure that patient safety is
not compromised, treatment eﬃcacy is documented, and technologic
improvements are integrated into the mainstream of HD therapy.

Dialysis prescription formulation

Standard prescription variables

The dialysis prescription is individually formulated at each treatment to

remove waste solutes and to normalize ﬂuid, electrolyte, and acid-base
balance to the greatest degree possible

. The prescription for a given

patient varies from session to session based on residual renal function, urine
production, biochemical status, and comorbid conditions. Variable compo-
nents of acute and chronic dialysis prescriptions are presented in

Table 1.

Sodium proﬁling

The capacity to proﬁle (or model) the dialysate sodium concentration to

accommodate the patient’s physiologic status better is now a standard
feature of delivery systems. Rapid solute removal and ultraﬁltration can
cause intradialytic osmotic and ﬂuid shifts that may lead to complications,
including hypovolemia, hypotension, cramping, nausea, vomiting, and
dialysis disequilibrium syndrome. Sodium proﬁling, in which the sodium
concentration of the dialysate is systematically altered during the dialysis
session, has been proposed as a means to lessen or prevent such signs
without promoting hypotension associated with constant low-sodium

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

prescriptions or positive sodium balance, postdialysis thirst, or interdialytic
weight gain associated with constant high-sodium prescriptions. To improve
the hemodynamic stability of animal patients predisposed to hypotension
and hypovolemia during dialysis, the delivery system can generate stepped
or linear adjustments in dialysate sodium that change the dialysate from
hypernatric (155–160 mmol/L) during the initial stages of the session to
hypo- or isonatric (140–150 mmol/L) at the end. The eﬃcacy of dialysate
sodium proﬁling has not been established conclusively but seems to beneﬁt
human patients predisposed to hemodynamic instability or excessive intra-
dialytic symptomology

[31–33]

. Linear high-to-low sodium proﬁling is

a standard HD prescription component at the University of California’s
HD units for cats and small dogs that are not hypertensive. Sodium proﬁling
seems to provide improved hemodynamic stability, although these obser-
vations need further validation.

Ultraﬁltration

As is done with dialysate sodium, the timing and rate of ultraﬁltration

can be modeled with modern delivery systems to match ﬂuid removal to
periods of greatest hemodynamic stability or support. Ultraﬁltration pro-
ﬁling can be coordinated with sodium proﬁling for each patient so that
maximal ﬂuid removal occurs when hypernatric dialysate proﬁles support
blood volume and vascular reﬁlling. In the absence of sodium proﬁling,
more eﬃcient and less symptomatic ﬂuid removal may be achieved by
increasing ultraﬁltration in the latter part the dialysis session. During this
time, solute clearance declines because the concentration gradient between

Table 1
Variable components of acute and chronic hemodialysis prescriptions

Variable

Acute prescription

Chronic prescription

Dialyzer type

Smaller surface area: 0.22–1.1 m

Larger surface area: 0.22–2.1 m

Blood ﬂow

Slower: 1–5 mL/kg/min

Faster: 10–25 mL/kg/min

Treatment length

Shorter: 2–4 hours

Longer: 4–6 hours

Dialysate composition

Sodium

Modeled

Constant: 145 or 150 mmol/L

Potassium

0 or 3 mmol/L

Bicarbonate

25–30 mmol/L

30–35 mmol/L

Phosphorus

Variable, usually 0

Other additives

Variable

Ultraﬁltration rate

Variable

Anticoagulation

Variable

Intradialytic medications

Mannitol

Likely; bolus followed by infusion Unlikely

Interdialytic interval

12–24 hours

48–96 hours

These variables are tailored at each treatment to correct a given patient’s ﬂuid, electrolyte,

and metabolic balance. This table compares guidelines for prescriptions commonly applied to
the ﬁrst few treatments (‘‘acute prescriptions’’) with those applied to later treatments (‘‘chronic
prescriptions’’).

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

blood and dialysate is reduced; thus, blood volume is less inﬂuenced by
associated osmotic ﬂuid shifts. Recommendations for dialysate and ultra-
ﬁltration proﬁling have not been established for animal dialysis but will be
developed as equipment with these capabilities becomes incorporated into
veterinary HD programs.

Staged azotemia reduction

Most animals initially presented for dialysis have marked azotemia, often

with blood urea nitrogen (BUN) in excess of 200 mg/dL, and consequently
have equally markedly elevated plasma osmolality

[34,35]

. Although it is

technically feasible to resolve even severe azotemia completely in a matter of
hours, decreases in azotemia that are too precipitous can result in dialysis
disequilibrium syndrome, the clinical manifestations of cerebral edema

[3,7,26]

. For this reason, the initial dialysis treatment for severely azotemic

animals (BUN [150) is deliberately prescribed to be ineﬃcient. Blood ﬂows
and treatment times are calculated to produce a reduction in the BUN that
is between 30% and 50% of the starting value but not to exceed a change of
100 mg/dL. The second treatment is usually prescribed to produce a 50% to
70% reduction in the BUN but is again limited to a 100-mg/dL decrease.
The BUN and creatinine can be safely normalized in most animals by the
third dialysis treatment.

In extremely small patients, decreasing azotemia slowly can be quite

challenging, even with the smallest dialyzers available and the lowest blood
ﬂows the machine permits; complete extraction of solutes and rapid
achievement of ﬁltration equilibrium occur as blood ﬂows through highly
eﬃcient dialyzers. In such patients, periods of ‘‘bypass’’ (during which blood
continues to ﬂow through the circuit and dialyzer but dialysate does not
circulate) can be interspersed with periods of active dialysis to permit
rebound of dialyzed solutes. This pattern allows extended gradual physio-
logic treatment with decreased risk of disequilibrium and increased time
over which to optimize ultraﬁltration. Reversing the attachments of the
blood lines to the catheter ports (connecting the withdrawal line to what
would normally be the return port and the return line to the usual
withdrawal port) decreases treatment eﬃcacy by increasing the blood
recirculation ratio

. Blood lines may be reversed as another measure

to intentionally render an initial treatment in a small patient less eﬃcient.

Single-needle techniques

Some dialysis delivery systems have software that permits treatment using

a single pathway for drawing and returning blood to the patient, termed
single-needle

treatment. In this situation, a Y-shaped piece connects the

arterial and venous lines of the extracorporeal circuit to the single catheter
hub. During the ﬁrst phase of the cycle, the venous line is clamped and the
pump pulls blood into the arterial line. The ﬁrst phase ends when (1) a preset

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

stroke volume is reached, (2) a preset pressure is reached, or (3) a preset time
elapses. These variables are individually manually adjustable but are
interdependent in nature. During the second phase of the cycle, the arterial
line is clamped, the venous line is released, and the pump returns blood
through the venous line. This cycle repeats for the duration of the treatment,
alternately drawing and returning blood to the patient. Adjustment of
stroke volume, pressure limit, or stroke duration modulates the rate of
dialysis delivery. Single-needle capabilities allow treatment of a patient that
has only one functioning catheter lumen in situations where replacement or
re-establishment of dual-lumen access is not a viable or safe option. Small
patients can be eﬀectively treated with standard extracorporeal circuitry;
larger patients require the addition of an expandable chamber to receive the
blood as it is drawn or a special circuit designed for single-needle treatment.
This treatment mode has reduced eﬃcacy compared with standard
treatment because of lower total blood ﬂow through the dialyzer and
because of recirculation of blood through the arterial line (ie, because of the
dead space in the Y-shaped piece and the catheter).

Management problems in chronic hemodialysis patients

HD extends longevity in dogs and cats with minimal intrinsic renal

function. Because these animals survive beyond the life expectancies that
conventional medical management provides, they demonstrate a spectrum of
pathologic and clinical conditions not typically recognized in conventionally
managed uremic animals. In addition, management of clinical conditions
that are frequently benignly neglected in animals with a natural progression
of disease becomes fundamental to successful management of the dialysis
patient. Malnutrition, speciﬁc amino acid depletion, hormonal derange-
ments, aluminum toxicosis, and metabolic bone disease can all produce
morbid consequences in dialysis-dependent chronically uremic animals and
thus can complicate routine and successful management of renal failure.

Uremia-related complications

Malnutrition

Malnutrition, one of the most pervasive complications of renal failure, is

accentuated in HD patients by nausea, vomiting and anorexia by direct
stimulation of the chemoreceptor trigger zone as well as by signiﬁcant oral
ulceration or gastrointestinal pathologic ﬁndings (

)

[37–39]

. Mucosal

lesions usually resolve within days of starting dialytic therapy, but most
animal HD patients remain inappetent or anorexic. This may be partially
a result of underdialysis and failure to remove small-molecular-weight solutes,
inadequate removal of middle-molecular weight substances (eg, leptin, which
has appetite-suppressive properties), or other comorbid conditions

[37]

HD patients exhibit increased energy and protein requirements at rest

and while undergoing treatment

. Resting energy expenditure in chronic

952

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

HD patients may be increased 10% to 20% over that of normal individuals
for ill-deﬁned reasons

[38,40]

. During HD treatments, exposure to bio-

incompatible dialyzer membranes as well as to foreign substances, such as
endotoxin in dialysate, may activate the complement system, leading to
systemic inﬂammation and increased protein catabolism. Production of
interleukin-1 and tumor necrosis factor-a, among other cytokines, may
induce degradation of muscle proteins and release of inappropriately high
amounts of amino acids

. Signiﬁcant amino acid loss during the dialysis

session has been documented in human beings and dogs

[41,42]

. Addition-

ally, metabolic acidosis, ubiquitous in renal failure patients, is a major
eﬀector of progressive muscle catabolism

In human beings, malnutrition has been correlated with increased

mortality and morbidity in patients with end-stage renal disease and
ARF, particularly during hospitalization

[39,44,45]

. Early and aggressive

nutritional intervention is therefore prudent in any dialysis patient.
Placement of an esophageal or gastric feeding tube or a dedicated catheter
for parenteral nutrition at the time of dialysis catheter placement enables
proactive nutritional management of HD patients’ improved protein-calorie
balance.

Hormonal derangements

Insulin resistance

Decreased anabolic capacity also plays a part in the malnutrition of HD

patients. Uremia renders target tissues, muscle tissue in particular, resistant
to select eﬀects of insulin, resulting in mild to moderate carbohydrate

MALNUTRITION

Decreased

appetite

Endocrinopathies

Dialysis membrane-

induced catabolism

Decreased

anabolism

Dialytic loss

of nutrients

Metabolic

acidosis

Protein-

restricted diet

Increased resting

energy expendature

Fig. 6. Factors contributing to malnutrition in hemodialysis (HD) patients. Malnutrition in
HD patients is a complex multifactorial phenomenon requiring ongoing assessment and
proactive global management.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

intolerance

; myocytes still retain the capacity to synthesize glycogen

normally but cannot perform normal glycolysis and glucose oxidation

In a 90% nephrectomy model, uremic rats demonstrated a 28% reduction in
total body glucose disposal compared with nonnephrectomized controls

. In a study using a euglycemic insulin clamp technique, chronically

uremic human subjects exhibited a 47% reduction in insulin-mediated
glucose metabolism when compared with control subjects

[48]

. The

hyperparathyroidism of chronic uremia has also been suggested to induce
carbohydrate intolerance by inhibition of insulin secretion by pancreatic b
cells

. Although this diminished capacity to use glucose may not be

severely debilitating in itself, it represents another detractor to the chronic
dialysis patient’s nutritional status.

Erythropoietin resistance

As renal function declines, production of erythropoietin ebbs, diminish-

ing the patient’s ability to stimulate red blood cell production in the bone
marrow, resulting in progressive nonregenerative anemia. Supplementation
with subcutaneous recombinant human erythropoietin (rHuEPO; Procrit;
Ortho Biotech, Bridgewater, NJ; EPOGEN, Amgen, Thousand Oaks, CA)
produces an initial erythroid response in most dogs and cats receiving short-
and long-term HD, which invariably require rHuEPO therapy

[50]

. Despite

the initial positive response, however, a pure red cell aplasia caused by anti-
rHuEPO antibodies develops in 20% to 70% of animal patients treated
chronically, rendering its continued use ineﬀective and contraindicated for
those patients

[50–53]

. Other confounding factors, such as iron deﬁciency,

ongoing blood loss, systemic inﬂammation, secondary hyperparathyroid-
ism, or hemolytic anemia, should be ruled out in animals with rHuEPO
resistance

[54,55]

. In human dialysis patients, carnitine supplementation can

improve response to rHuEPO

[53]

Darbepoetin alfa (Aranesp, Amgen) is a novel erythropoietic peptide that

has been proven to be as eﬀective as rHuEPO in stimulating red blood cell
production in human patients with chronic renal failure

[53,56,57]

. Com-

pared with rHuEPO, darbepoetin has a longer half-life and greater potency,
enabling clinical eﬃcacy with less frequent administration

[53,56,57]

. In

a limited number of veterinary patients, darbepoetin has produced an
eﬀective increase in hematocrit with the advantage of less frequent injections
than rHuEPO requires. Darbepoetin’s increased half-life and potency may
result in decreased human antigen exposure to animal patients compared
with rHuEPO. In theory, this may result in a lower rate of antibody
development in animal patients, making darbepoetin more eﬀective and safer
than rHuEPO for erythropoietin replacement in dialysis patients.

Aluminum toxicosis

Chronic use of high doses of aluminum-based phosphate binders can lead

to aluminum toxicosis in small animals. Because excess aluminum is chieﬂy

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

excreted in the urine, animals in renal failure may have marked exposure to
aluminum from phosphate binders. Additionally, renal failure patients have
increased intestinal absorption of aluminum as well as diminished aluminum
excretory capacity

[58–60]

. HD patients have an additional risk of peracute

toxicosis if the water used for dialysate preparation has elevated aluminum
levels

[61]

. Clinical signs of aluminum toxicosis in small animals are usually

neurologic or neuromuscular and may be subtle (eg, mild weakness or
exercise intolerance, mental dullness) or profound (eg, severe obtundation,
coma, paresis). Microcytosis and hypochromia, usually with anemia, often
also develop and can precede neurologic signs

[62]

. Treatment of aluminum

toxicosis with discontinuation of aluminum-based phosphate binders and
aluminum chelation with deferoxamine has been successful in human
patients

[63]

and also at the University of California at Davis dialysis unit

in several patients with severe clinical signs of aluminum toxicosis after
chronic administration of high (up to 500 mg/kg/d) doses of aluminum
hydroxide. Because most chronic HD patients have mild to moderate
hypercalcemia, use of calcium-based phosphate binders is problematic. A
new nonaluminum- and noncalcium-based phosphate binder, sevelamer
hydrochloride (Renagel, Genzyme Corp., Cambridge, MA) is available and
performs well in human patients; however, this agent is costly and has not
been proven eﬃcacious or been well studied in animals

[64,65]

Metabolic bone disease

Secondary hyperparathyroidism develops to some degree in most animals

with end-stage renal disease secondary to phosphorus retention; osteopenia
resulting from chronic inappropriate calcium mobilization may occur as
a sequela

[66–68]

. Although this rarely manifests clinically in conventionally

managed uremic animals

[66,68]

, chronically dialyzed patients may be at

risk for pathologic fractures as a result of bone demineralization. Concur-
rent administration of aluminum hydroxide may place these animals at
further risk of skeletal pathologic conditions because of aluminum
accumulation in bone

[69,70]

. Calcitriol or other similar therapy is the

standard of care in human renal failure patients and is used variably in
animal patients to help prevent or reverse hyperparathyroidism and its
sequelae. Such therapy is often contraindicated in canine and feline HD
patients, however, because of preexisting hypercalcemia. Newer vitamin D
derivatives (eg, 22-oxacalcitriol, paricalcitol, doxercalciferol) may provide
therapeutic options to control parathyroid hormone secretion in animals
without exacerbating hypercalcemia

[71–73]

. Even more promising for HD

patients are newer calcimimetic agents (eg, cinacalcet hydrochloride) that
control parathyroid hormone secretion by increasing sensitivity of the
calcium-sensing membrane receptor on parathyroid cells, often actually
reducing serum calcium concentration

[71,74–76]

. Although 22-oxacalcitriol

has been studied in research dogs

[73]

, clinical trials with calcimimetics have

not yet been conducted in animals.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Dialysis-related complications

Carnitine and taurine deﬁciencies

Carnitine deﬁciency occurs in chronically dialyzed patients as a result of

decreased protein intake, decreased protein synthesis, and direct loss of
carnitine through the dialyzer membrane

[77,78]

. Dialysis-induced carnitine

deﬁciency in people is associated with cardiomyopathy, muscle weakness,
intradialytic hypotension, and erythropoietin-resistant anemia, all of which
are ameliorated by carnitine supplementation

[79–82]

. Taurine is also lost

through dialyzer membranes

[83]

. Given that carnitine deﬁciency–associated

cardiomyopathy has been identiﬁed in dogs and taurine deﬁciency is
associated with cardiomyopathy in dogs and cats and retinopathy in cats,
supplementation of carnitine and taurine in chronic (>1 month) animal
dialysis patients seems well justiﬁed

[83–88]

Dialysis catheter dysfunction

The most common complications seen with HD catheters are diminished

eﬀective blood ﬂow rates and bacterial infection of the catheter

[89,90]

. Blood

ﬂow rates may decrease because of thrombus formation within the catheter
lumen or at the portals, thrombotic or stenotic changes within the vessel itself,
or ﬁbrin sheath formation around the catheter

[5,6,90]

. With chronic use,

venous stenosis can occur distal to the catheter tip, causing diminished blood
ﬂow or an unacceptably high recirculation ratio (ﬂow of processed blood from
the outﬂow portal directly back to the inﬂow portal)

[6,91]

. An acute rise in

recirculation ratio strongly suggests venous thrombosis or stenosis formation,
which can be conﬁrmed with angiography and ﬂuoroscopy

[6,90,91]

. A

thrombus at the catheter tip often can be dislodged with forceful saline
injection. Failing that, thrombolytic agents, such as streptokinase or tissue-
type plasminogen activator, can be infused through the catheter, or a per-
cutaneous thrombectomy technique involving maceration and aspiration of
the clot fragments (AngioJet System, Possis Medical, Minneapolis, Minne-
sota) may be employed

[89,92]

. These procedures are costly and may carry risk

to the patient (eg, the possibility of uncontrollable hemorrhage with throm-
bolytic agents); thus, the absolute need to preserve a given vascular access
must be weighed carefully. Clot formation within one or both catheter lumens
can occur during the interdialytic interval, despite the fact that the catheters
are routinely locked with heparin and patients are maintained on antithrom-
botic doses of aspirin. Clots within the catheter can usually be dislodged with
aspiration or saline ﬂushes or disrupted mechanically with a guidewire passed
down the catheter

[89,93]

. Venous stenosis usually mandates replacement of

the catheter, either in the contralateral jugular vein if the cranial cava is
unaﬀected or in the same vein with the vascular portals placed beyond the
stenotic region. Balloon angioplasty or placement of a stent graft in a stric-
tured or stenotic vascular segment is commonly performed in human patients
but has not been investigated in veterinary patients for this purpose

[5,94]

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Human studies have demonstrated ﬁbrin sheath formation around

indwelling central venous catheters as early as 48 hours after placement

[5,6]

. These thin ﬂexible sheaths extend from the venotomy site along the

length of the catheter like a plastic bag, enveloping the portals and acting like
a one-way valve that permits outﬂow of blood but precludes inﬂow. Fibrin
sheaths are seen in veterinary HD patients and usually manifest clinically as
decreased ﬂow or as an inability to aspirate from the arterial or both catheter
ports. Sheaths can be removed via pharmacologic lysis or mechanical
stripping

[5,6,90,93,95]

. In human beings, ﬁbrin sheath stripping has greater

than 90% eﬃcacy for normalizing catheter ﬂow rates

. In the stripping

procedure, a wire snare introduced into the femoral vein is threaded up to the
catheter, guided to encircle the catheter base at the venotomy site, and then
tightened around the catheter. Drawing the snare oﬀ the catheter strips the
ﬁbrin sheath, which usually is embolized asymptomatically

. The presence

of a ﬁbrin sheath as the cause of diminished ﬂow should be ruled out before
guidewire exchange of a catheter so as to avoid the possibility of placing
a new catheter directly into the extant sheath. Sheath stripping is a feasible
but untried method of catheter salvage in animals.

Eﬀective catheter ﬂow may dramatically decrease without any apparent

impact on metered blood ﬂow rates if increased recirculation ratios are
present

[6,91]

. Recirculation occurs when conditions favor the direct intake

into the catheter’s withdrawal port of just-puriﬁed blood returning from the
dialyzer. Under normal conditions, staggering and spacing of the intake and
outﬂow ports of the dialysis catheter as well as placement of the outﬂow port
in a ‘‘downstream’’ vascular position relative to the intake port prevent
signiﬁcant recirculation from occurring. Additionally, high-volume blood
ﬂow in the central vascular segments accessed for HD (eg, cranial vena cava,
right atrium) quickly dilutes, returning puriﬁed blood with waste-laden
blood from peripheral capillary beds. Development of a stenosis or
thrombosis near the venous outﬂow disrupts normal blood ﬂow patterns
and pools returning blood, rendering it available for reuptake in the arterial
port

. Reprocessing of puriﬁed blood before its passage through peripheral

capillary beds diminishes the amount of waste solute available for removal
without diminishing the total amount of waste solute in the body. An HD
treatment that produces substantially less urea reduction than expected
based on blood volume processed suggests the presence of recirculation

[91]

Recirculation ratios can be quantiﬁed in the clinical setting by several means,
including ultrasound or infrared measurement of saline dilution, thermal
dilution, or ionic dialysance methods

[6,28,96]

. Tolerable degrees of blood

recirculation should be below 10% to 15% and ideally below 3% to 7%

Bacterial catheter infection

Bacterial catheter infection is a well-recognized and often devastating

complication of transcutaneous catheter use, necessitating protracted
antibiotic therapy and potentially resulting in temporary or permanent loss

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

of vascular access, bacteremia, bacterial endocarditis, and the death of the
patient

[5,16]

. In human beings, use of catheters for HD access is an

important risk factor for development of infections, the second most
common cause of death in HD patients, and a major contributor to HD-
associated morbidity and mortality

[11,13,23,24,97]

. Catheter infection

should be suspected when the catheter exit site is erythematous or indurated
or when pus is present, if a new cardiac murmur is detected, if fever
(temperature [101(F in a signiﬁcantly azotemic patient) is present with no
other obvious cause, if a patient develops ‘‘chills’’ during dialysis treatment,
or if the body temperature rises signiﬁcantly after initiation of dialysis

[5,13,98]

. In these situations, swabs of the catheter exit site, the heparin locks,

or peripheral blood should be submitted for aerobic and anaerobic culture.
One human study documented the presence of exit site infection in 46% of
temporary dialysis catheter–associated bacteremias

[99]

. In this report, the

likelihood of bacteremia occurring within 1 day of clinical evidence of exit
site infection was 1.9%

[99]

. Incidence of bacteremia increased to 13.4% by

the second day of exit site infection, emphasizing the need for fastidious
catheter care and proactive management of suspicious catheter sites

[16,99]

If a positive bacterial culture is obtained from the exit site but cultures of

the blood or heparin locks are negative, appropriate antibiotic treatment
should be instituted while the catheter remains in place. When positive
bacterial culture of the heparin locks or the blood conﬁrms catheter
infection, the question of whether to remove the dialysis catheter must be
confronted. Marr et al

[98]

evaluated the eﬃcacy of catheter salvage

procedures in the face of bacteremia and demonstrated a failure rate of
68% even with prolonged antibiotic therapy. Ideally, catheter removal
followed by intensive antibiotic therapy and demonstration of negative
blood cultures should precede replacement of the dialysis catheter

[5,16,97,98]

. Unfortunately, most dialysis-dependent animals cannot live

asymptomatically for more than 3 to 5 days oﬀ dialysis, necessitating re-
establishment of vascular access within this interval. Human studies
examining the cost-eﬀectiveness of infected catheter salvage compared
antibiotic treatment with the catheter left in place, guidewire exchange of
the catheter, and catheter removal and replacement

[100,101]

. Guidewire

exchange of the catheter provided signiﬁcant cost savings over other
management strategies

[100,101]

. Despite the high odds of failure, catheter

salvage (or vessel/access site salvage with guidewire exchange of the catheter
at the least) must be strongly considered in companion animals if the
infection does not seem to be life threatening, because alternate sites and
methods of vascular access are severely limited.

Several protocols have been proposed for treatment of bacterial catheter

infection using high concentrations of antibiotic with heparin- or tauroli-
dine-based catheter lock solutions designed to destroy the luminal bioﬁlm
that permits bacterial colonization and persistence of infection. Several
recent in vitro studies have evaluated the stability or in vitro eﬃcacy of

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

several antibiotic/heparin catheter lock solutions

[102–104]

. Vercaigne et al

[104]

ﬁlled central venous catheters with gentamicin, cefazolin, ceftazidime,

or vancomycin at a concentration of 10 mg/mL and showed that signiﬁcant
antibiotic activity (

5 mg/mL) persisted within the catheter lumens after

incubation at 37(C for 72 hours. Droste et al

[103]

combined ciproﬂoxacin,

ﬂucloxacillin, linezolid, ciproﬂoxacin, and a teicoplanin-ciproﬂoxacin com-
bination with varying concentrations of heparin and found that use of
higher heparin concentrations (3500–10,000 U/mL) produced more stable
solutions. These solutions showed good eﬀects against the speciﬁcally
targeted pathogens, including vancomycin-resistant enterococci (linezolid-
heparin) and Pseudomonas aeruginosa (ciproﬂoxacin-heparin)

[103]

. Krish-

nasami et al

[102]

reported successful catheter salvage in 50% of human

dialysis patients with antibiotic/heparin locks and systemic antibiotics.

Hemodialysis therapy in small animal practice

Change in the etiology of severe acute uremia since 1996

Since 1996, the causes of severe acute uremia in dogs presented for HD to

the University of California at Davis have shifted from most with
nephrotoxicosis (60%) and fewer with infectious causes (15%) to an even
repartition of toxic and infectious causes (40% each)

. This change is

mostly a result of the re-emergence of leptospirosis, which is now the leading
cause of acute uremia in dogs (40%) in California, followed by ethylene
glycol toxicosis (30%)

[105]

Etiology of acute uremia in cats presented to the University of California

at Davis for dialytic therapy have also shifted. In 1997, Langston et al

reported etiology and outcomes for the 29 cats treated with HD at the
University of California at Davis from 1993 through 1996. Of these cats,
60% were presented for ethylene glycol toxicosis. In recent years, acute
ureteral obstruction has emerged as the leading cause of acute uremia in cats

. Since 1996, more than 120 cats have been dialyzed at the University of

California at Davis and at the University of California Veterinary Medical
Center in San Diego. Of these cats, approximately 45% were presented for
uremia related to ureteral obstruction compared with 20% for nephrotox-
icosis (V. Pantaleo, DVM, J.R. Fischer, DVM, unpublished data, 2003). Of
the nephrotoxicosis cases, roughly 60% of cats had ingested ethylene glycol
and 40% had ingested lily (V. Pantaleo, DVM, J.R. Fischer, DVM,
unpublished data, 2003).

Outcomes and prognosis

The prognosis for recovery from acute uremia in dogs or cats provided

HD depends on the etiology, extent of renal damage, comorbid diseases,
and presence of multiple organ system involvement. There is little
documentation in the veterinary literature to predict the importance of

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

these independent variables accurately. A recent review of 138 dogs with
severe acute uremia requiring HD after failure to respond to conventional
medical management revealed a survival rate of nearly 40%

. During the

ﬁrst 6 years of the HD program at the University of California at Davis, 52
dogs were treated for acute uremia with a global survival rate of 30%.
Survival from infectious (60%), hemodynamic, and metabolic causes (40%)
was greater than survival from toxic causes (20%)

. During this same

period, of the 23 acutely uremic (ARF and acute exacerbation of chronic
renal failure) cats in Langston et al’s report

, global survival was 43%.

Surprisingly, in this report, survival for cats with toxic causes (chieﬂy
ethylene glycol) approached 60%

Since 1996, for 86 dogs with severe acute uremia treated with HD, global

survival increased to 50%. This improvement is mostly a result of the
increased incidence of infectious etiologies from 15% to 40% and the re-
emergence of leptospirosis. The outcome for dogs with acute leptospirosis is
particularly favorable, with an 85% survival rate with either severe (dialysis-
dependent) or milder forms (medically manageable) of ARF

[34,106]

Survival of dogs with severe acute uremia requiring HD was independent
of the degree of azotemia at presentation and was directly inﬂuenced by the
underlying etiology

. For most surviving dogs, renal function recovered

substantially by the time of discharge and progression to chronic renal
failure was observed only rarely. Global survival for cats requiring HD has
improved to 56% since 1996 (J.R. Fischer, DVM, unpublished data, 2003).
The increased survival rate is largely a result of the dramatic increase in the
number of cats presented for acute ureteral obstruction

. Since 1996,

35% of the cats presented for HD have had acute ureteral obstruction, and
of these cats, more than 70% survived (V. Pantaleo, DVM, unpublished
data, 2003). Overall, HD substantially increases the global survival for dogs
and cats with severe acute uremia beyond what would be expected with
conventional management.

Summary

HD can be a life-saving intervention for dogs and cats with severe acute

uremia. The metabolic stability provided by this treatment modality can
aﬀord the clinician time to diagnose the underlying etiology of the renal
dysfunction and thus provide clients with improved prognostic data. HD
can also extend the life of patients with end-stage renal disease that cannot
be adequately managed with conventional means, giving clients time to
adjust to a terminal diagnosis, to prepare for chronic dialysis therapy, or to
condition a pet for renal transplantation. Dialysis can also eﬀectively
manage refractory states of volume overload and can expediently remove
many toxins and pharmacologic agents from the bloodstream. Veterinary
HD referral centers are currently available in California, New York,
Massachusetts, and Pennsylvania.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Appendix A: North American veterinary facilities oﬀering acute and chronic
hemodialysis services

The Animal Medical Center
510 E. 62nd Street
New York, NY 10021
Main: (212) 838-8100
Dialysis room: (212)329-8618
e-mail address:

hemodialysis@amcny.org

Dr. Kathy Langston

Tufts Foster Hospital for Small Animals
200 Westboro Road
North Grafton, MA 01536
(508) 839-5395 ext. 84681
Dr. Mary Labato
Dr. Linda Ross

University of California Veterinary Medical Teaching Hospital
Companion Animal Hemodialysis Unit
1 Garrod Drive
Davis, CA 95616
(530) 752-1393
Dr. Thierry Francey
Dr. Larry Cowgill

University of California Veterinary Medical Center at San Diego
Renal Medicine/Hemodialysis Service
PO Box 9415
6525 Calle del Nido
Rancho Santa Fe, CA 92067
(858) 759-7235
Dr. Julie Fischer
Dr. Larry Cowgill

Veterinary Hospital of the University of Pennsylvania
Hemodialysis Center
Matthew J. Ryan Veterinary Hospital
3900 Delancey Street
University of Pennsylvania
Philadelphia, PA 19104
(215) 898-4680
Dr. Reid Groman

Appendix B: Hemodialysis referral guidelines for practitioners

Call early and refer early: the clinicians serving HD centers can guide

selection of patients likely to beneﬁt from dialytic intervention and oﬀer

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

input on medical management of uremic animals that may not require HD.
In general, patient stability and odds of a successful outcome with HD
decrease the longer the uremia and its attendant metabolic and ﬂuid
derangements persist. Referral for dialytic therapy is often life saving for
acutely uremic patients that do not respond to appropriate and aggressive
medical management within 12 to 24 hours, and referral for dialysis is
a logistic option for the clients.

Spare the jugular veins: the blood ﬂow rates required to perform HD

mandate placement of a large-gauge jugular catheter with portals ideally
situated in the right atrium or cranial vena cava. The condition of the
jugular veins on presentation often determines percutaneous catheter versus
surgical catheter placement and use of local versus general anesthesia.
Additionally, prior trauma to the vein increases the odds of placement-
related complications, such as venous tearing, regardless of placement
method. For these reasons, jugular venipuncture and jugular catheters
should be avoided in patients in which HD remains an option. If jugular
vein access is unavoidable, use of the left jugular vein is preferred (sparing
the right jugular vein completely) and adequate hemostasis after venipunc-
ture is essential to avoid hematoma formation.

Fully inform clients before referral: HD is an emotionally and ﬁnancially

intensive therapy with no hard guarantees of a successful outcome. It
involves deﬁned risk to an already compromised and often unstable patient
and usually requires sequential or alternate-day treatments over weeks to
months. HD is an outstanding bridging mechanism that often permits life-
saving repair of renal injury in patients when no other therapeutic options
exist, but clients must understand that dialysis does not ‘‘ﬁx’’ damaged
kidneys. Usually, it is impossible to determine at the outset how long
therapy must continue to allow a patient’s tubular function to resume. In
general, with severe acute tubular necrosis, clients should be ﬁnancially and
emotionally prepared to undertake 2 to 4 weeks of dialytic therapy; 7 to 14
days is the earliest reasonable window during which to expect signs of
resumption of renal function, although some patients can recover more
quickly. Conversely, some patients have recovered renal function only after
many months of dialysis dependency. Prognosis and duration of therapy
vary tremendously from patient to patient and depend on the cause and
degree of renal insult as well as on patient condition and comorbidities.
Dialysis referral centers can often provide written and verbal information to
clients before referral to ensure that they are fully and accurately educated
regarding the advantages and limitations of dialytic intervention.

References

[1] Daugirdas JT, Van Stone J. Physiologic principles and urea kinetic modelling. In:

Daugirdas JT, Blake PG, Ing TS, editors. Handbook of dialysis. Philadelphia: Lippincott
Williams & Wilkins; 2001. p. 15–45.

962

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

[2] Cowgill L, Langston C. Role of hemodialysis in the management of dogs and cats with

renal failure. In: Polzin DJ, editor. Renal dysfunction. vol. 26. Philadelphia: Harcourt
Brace & Company; 1996. p. 1347–78.

[3] Schulman G, Himmelfarb J. Hemodialysis. In: Brenner BM, editor. The kidney. vol. 2.

Philadelphia: WB Saunders; 2004. p. 2564–624.

[4] Daugirdas JT, Van Stone JC, Boag JT. Hemodialysis apparatus. In: Daugirdas JT, Blake

PG, Ing TS, editors. Handbook of dialysis. 3rd edition. Philadelphia: Lippincott Williams
& Wilkins; 2001. p. 46–58.

[5] Schwab SJ, Beathard G. The hemodialysis catheter conundrum: hate living with them,

but can’t live without them. Kidney Int 1999;56(1):1–17.

[6] Leblanc M, Bosc JY, Paganini EP, Canaud B. Central venous dialysis catheter

dysfunction. Adv Ren Replace Ther 1997;4(4):377–89.

[7] Cowgill L, Elliott D. Hemodialysis. In: DiBartola SP, editor. Fluid therapy in small

animal practice. Philadelphia: WB Saunders; 2000. p. 528–47.

[8] Langston CE, Cowgill LD, Spano JA. Applications and outcome of hemodialysis in cats:

a review of 29 cases. J Vet Intern Med 1997;11(6):348–55.

[9] Sternbach G, Seldinger SI. Catheter introduction on a ﬂexible leader. J Emerg Med 1990;

8(5):635–7.

[10] Richard HM III, Hastings GS, Boyd-Kranis RL, Murthy R, Radack DM, Santilli JG,

et al. A randomized, prospective evaluation of the Tesio, Ash split, and Opti-ﬂow
hemodialysis catheters. J Vasc Interv Radiol 2001;12(4):431–5.

[11] Gulati S, Sahu KM, Avula S, Sharma RK, Ayyagiri A, Pandey CM. Role of vascular

access as a risk factor for infections in hemodialysis. Ren Fail 2003;25(6):967–73.

[12] National Kidney Foundation. K/DOQI clinical practice guidelines for vascular access,

2000. Am J Kidney Dis 2001;37(Suppl):S137–81.

[13] Allon M, Depner TA, Radeva M, Bailey J, Beddhu S, Butterly D, et al. Impact of dialysis

dose and membrane on infection-related hospitalization and death: results of the HEMO
Study. J Am Soc Nephrol 2003;14(7):1863–70.

[14] US Renal Data System. USRDS 2003 annual data report: atlas of end-stage renal disease

in the United States. Bethesda: National Institutes of Health, National Institute of
Diabetes and Digestive and Kidney Diseases; 2003.

[15] Ifudu O, Macey LJ, Homel P, Hyppolite JC, Hong J, Sumrani N, et al. Determinants of

type of initial hemodialysis vascular access. Am J Nephrol 1997;17(5):425–7.

[16] Nassar GM, Ayus JC. Infectious complications of the hemodialysis access. Kidney Int

2001;60(1):1–13.

[17] Adin CA, Gregory CR, Adin DB, Cowgill LD, Kyles AE. Evaluation of three

peripheral arteriovenous ﬁstulas for hemodialysis access in dogs. Vet Surg 2002;31(5):
405–11.

[18] Rosenblatt M, Stainken B, Weiss M, Carid IJ. LifeSite totally implanted HD system

versus Tesio Cath: results of a comparative trial [abstract]. J Vasc Interv Radiol 2002;13(2
Suppl):S39.

[19] Schwab SJ, Weiss MA, Rushton F, Ross JP, Jackson J, Kapoian T, et al. Multicenter

clinical trial results with the LifeSite hemodialysis access system. Kidney Int 2002;62(3):
1026–33.

[20] Stas KJ, Vanwalleghem J, De Moor B, Keuleers H. Trisodium citrate 30% vs. heparin

5% as catheter lock in the interdialytic period in twin- or double-lumen dialysis catheters
for intermittent haemodialysis. Nephrol Dial Transplant 2001;16(7):1521–2.

[21] Weijmer MC, Debets-Ossenkopp YJ, Van De Vondervoort FJ, ter Wee PM. Superior

antimicrobial activity of trisodium citrate over heparin for catheter locking. Nephrol Dial
Transplant 2002;17(12):2189–95.

[22] Ash SR, Mankus RA, Sutton JM, et al. Concentrated sodium citrate (23%) for catheter

lock. Hemodial Int 2000;4:22–31.

[23] Dasgupta MK. Bioﬁlms and infection in dialysis patients. Semin Dial 2002;15(5):338–46.

963

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

[24] Allon M. Prophylaxis against dialysis catheter-related bacteremia with a novel

antimicrobial lock solution. Clin Infect Dis 2003;36(12):1539–44.

[25] Quarello F, Forneris G. Prevention of hemodialysis catheter-related bloodstream

infection using an antimicrobial lock. Blood Purif 2002;20(1):87–92.

[26] Cowgill LD, Langston CE. Role of hemodialysis in the management of dogs and cats with

renal failure. Vet Clin North Am Small Anim Pract 1996;26(6):1347–78.

[27] Ward RA. Ultrapure dialysate: a desirable and achievable goal for routine hemodialysis.

Semin Dial 2000;13(6):378–80.

[28] Petitclerc T. Festschrift for Professor Claude Jacobs. Recent developments in

conductivity monitoring of haemodialysis session. Nephrol Dial Transplant 1999;
14(11):2607–13.

[29] Polaschegg HD. Automatic, noninvasive intradialytic clearance measurement. Int J Artif

Organs 1993;16(4):185–91.

[30] Kuhlmann U, Goldau R, Samadi N, Graf T, Gross M, Orlandini G, et al. Accuracy and

safety of online clearance monitoring based on conductivity variation. Nephrol Dial
Transplant 2001;16(5):1053–8.

[31] Raja RM. Sodium proﬁling in elderly haemodialysis patients. Nephrol Dial Transplant

1996;11(Suppl 8):42–5.

[32] Coli L, Ursino M, Donati G, Cianciolo G, Soverini ML, Baraldi O, et al. Clinical

application of sodium proﬁling in the treatment of intradialytic hypotension. Int J Artif
Organs 2003;26(8):715–22.

[33] Gerrish M, Little J. The eﬀect of proﬁling dialysate sodium and ultraﬁltration on patient

comfort and cardiovascular stability during haemodialysis. EDTNA ERCA J 2003;29(2):
61, 65–6, 68–70.

[34] Francey T, Cowgill LD. Use of hemodialysis for the management of acute renal failure in

the dog: 124 cases (1990–2001) [abstract]. Presented at the 20th Annual American College
of Veterinary Internal Medicine Forum. Dallas, Texas, May 29–June 1, 2004.

[35] Pantaleo V, Francey T, Fischer JR, Cowgill LD. Application of hemodialysis for the

management of acute uremia in cats: 119 cases (1993–2003) [abstract accepted]. Presented
at the 22nd Annual American College of Veterinary Internal Medicine Forum.
Minneapolis, Minnestota, June 9–12, 2004.

[36] Level C, Lasseur C, Chauveau P, Bonarek H, Perrault L, Combe C. Performance of twin

central venous catheters: inﬂuence of the inversion of inlet and outlet on recirculation.
Blood Purif 2002;20(2):182–8.

[37] Chazot C, Laurent G, Charra B, Blanc C, VoVan C, Jean G, et al. Malnutrition in long-

term haemodialysis survivors. Nephrol Dial Transplant 2001;16(1):61–9.

[38] Ikizler TA, Wingard RL, Sun M, Harvell J, Parker RA, Hakim RM. Increased energy

expenditure in hemodialysis patients. J Am Soc Nephrol 1996;7(12):2646–53.

[39] Hakim RM, Levin N. Malnutrition in hemodialysis patients. Am J Kidney Dis 1993;

21(2):125–37.

[40] Neyra R, Chen KY, Sun M, Shyr Y, Hakim RM, Ikizler TA. Increased resting energy expen-

diture in patients with end-stage renal disease. J Parenter Enteral Nutr 2003;27(1):36–42.

[41] Ikizler TA, Flakoll PJ, Parker RA, Hakim RM. Amino acid and albumin losses during

hemodialysis. Kidney Int 1994;46(3):830–7.

[42] Elliott DA, Marks SL, Cowgill LD, Kass PH, Rogers QR. Eﬀect of hemodialysis on

plasma amino acid concentrations in healthy dogs. Am J Vet Res 2000;61(8):869–73.

[43] Mitch W. Inﬂuence of metabolic acidosis on nutrition. Am J Kidney Dis 1997;29:

xlvii–viii.

[44] Druml W. Nutritional management of acute renal failure. Am J Kidney Dis 2001;

37(1 Suppl2):S89.

[45] Ikizler TA, Greene JH, Yenicesu M, Schulman G, Wingard RL, Hakim RM.

Nitrogen balance in hospitalized chronic hemodialysis patients. Kidney Int Suppl 1996;
57:S53–6.

964

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

[46] DeFronzo RA, Smith D, Alvestrand A. Insulin action in uremia. Kidney Int Suppl 1983;

16:S102–14.

[47] Stein PP, Hunt WA, Johnson CM, DeFronzo RA, Smith JD. Insulin resistance in uremia:

an in vivo and in vitro study. Metabolism 1989;38(6):562–7.

[48] DeFronzo RA, Alvestrand A, Smith D, Hendler R, Hendler E, Wahren J. Insulin

resistance in uremia. J Clin Invest 1981;67(2):563–8.

[49] Mak RH, Bettinelli A, Turner C, Haycock GB, Chantler C. The inﬂuence of

hyperparathyroidism on glucose metabolism in uremia. J Clin Endocrinol Metab 1985;
60(2):229–33.

[50] Cowgill LD, James KM, Levy JK, Browne JK, Miller A, Lobingier RT, et al. Use of

recombinant human erythropoietin for management of anemia in dogs and cats with
renal failure. J Am Vet Med Assoc 1998;212(4):521–8.

[51] Langston C, Reine N, Kittrell D. The use of erythropoietin. Veterinary Clin Small Anim

Pract 2003;33:1245–60.

[52] Brassard JA, Meyers KM, Person M, Dhein CR. Experimentally induced renal failure in

the dog as an animal model of uremic bleeding. J Lab Clin Med 1994;124(1):48–54.

[53] Cowgill LD, Neal L, Egrie JC, Browne JK, Miller A. Antigenicity of recombinant human

erythropoietin (r-HuEPO) in normal dogs [abstract]. J Vet Intern Med 1994;8:167.

[54] Massry SG. Pathogenesis of the anemia of uremia: role of secondary hyperparathyroid-

ism. Kidney Int Suppl 1983;16:S204–7.

[55] King LG, Giger U, Diserens D, Nagode LA. Anemia of chronic renal failure in dogs.

J Vet Intern Med 1992;6(5):264–70.

[56] Vanrenterghem Y, Barany P, Mann JF, Kerr PG, Wilson J, Baker NF, et al. Randomized

trial of darbepoetin alfa for treatment of renal anemia at a reduced dose frequency
compared with rHuEPO in dialysis patients. Kidney Int 2002;62(6):2167–75.

[57] Ibbotson T, Goa KL. Darbepoetin alfa. Drugs 2001;61(14):2097–104; discussion 2105–6.
[58] Drueke TB. Intestinal absorption of aluminium in renal failure. Nephrol Dial Transplant

2002;17(Suppl 2):13–6.

[59] Cannata-Andia JB, Fernandez-Martin JL. The clinical impact of aluminium overload in

renal failure. Nephrol Dial Transplant 2002;17(Suppl 2):9–12.

[60] Ittel TH, Kluge R, Sieberth HG. Enhanced gastrointestinal absorption of aluminium

in uraemia: time course and eﬀect of vitamin D. Nephrol Dial Transplant 1988;3(5):
617–23.

[61] de Wolﬀ FA, Berend K, van der Voet GB. Subacute fatal aluminum poisoning in dialyzed

patients: post-mortem toxicological ﬁndings. Forensic Sci Int 2002;128(1–2):41–3.

[62] Mahieu S, del Carmen Contini M, Gonzalez M, Millen N, Elias MM. Aluminum toxicity.

Hematological eﬀects. Toxicol Lett 2000;111(3):235–42.

[63] Ogborn MR, Dorcas VC, Crocker JF. Deferoxamine and aluminum clearance in pediatric

hemodialysis patients. Pediatr Nephrol 1991;5(1):62–4.

[64] Loghman-Adham M. Safety of new phosphate binders for chronic renal failure. Drug Saf

2003;26(15):1093–115.

[65] Cizman B. Hyperphosphataemia and treatment with sevelamer in haemodialysis patients.

Nephrol Dial Transplant 2003;18(Suppl 5):v47–9.

[66] Nagode LA, Chew DJ, Podell M. Beneﬁts of calcitriol therapy and serum phosphorus

control in dogs and cats with chronic renal failure. Both are essential to prevent of suppress
toxic hyperparathyroidism. Vet Clin North Am Small Anim Pract 1996;26(6):1293–330.

[67] Polzin DJ, Osborne CA, Jacob F, Ross S. Chronic renal failure. In: Ettinger SJ, Feldman

E. 5th edition, editors. Textbook of veterinary internal medicine. vol. 2. Philadelphia: WB
Saunders; 2000p. 1634–62.

[68] Barber PJ, Elliott J. Feline chronic renal failure: calcium homeostasis in 80 cases

diagnosed between 1992 and 1995. J Small Anim Pract 1998;39(3):108–16.

[69] Malluche HH. Aluminium and bone disease in chronic renal failure. Nephrol Dial

Transplant 2002;17(Suppl 2):21–4.

965

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

[70] Malluche HH, Faugere MC, Friedler RM, Matthews C, Fanti P. Calcitriol, parathyroid

hormone, and accumulation of aluminum in bone in dogs with renal failure. J Clin Invest
1987;79(3):754–61.

[71] Drueke TB. Treatment of secondary hyperparathyroidism with vitamin D derivatives and

calcimimetics before and after start of dialysis. Nephrol Dial Transplant 2002;17(Suppl
11):20–2.

[72] Martin KJ, Gonzalez EA. Vitamin D analogues for the management of secondary

hyperparathyroidism. Am J Kidney Dis 2001;38(5 Suppl 5):S34–40.

[73] Monier-Faugere MC, Geng Z, Friedler RM, Qi Q, Kubodera N, Slatopolsky E, et al.

22-oxacalcitriol suppresses secondary hyperparathyroidism without inducing low bone
turnover in dogs with renal failure. Kidney Int 1999;55(3):821–32.

[74] Urena P. Use of calcimimetics in uremic patients with secondary hyperparathyroidism:

review. Artif Organs 2003;27(9):759–64.

[75] Urena P, Frazao JM. Calcimimetic agents: review and perspectives. Kidney Int Suppl

2003;(85):S91–6.

[76] Bardin T. Musculoskeletal manifestations of chronic renal failure. Curr Opin Rheumatol

2003;15(1):48–54.

[77] Matera M, Bellinghieri G, Costantino G, Santoro D, Calvani M, Savica V. History of

L-carnitine: implications for renal disease. J Ren Nutr 2003;13(1):2–14.

[78] Evans AM, Fornasini G. Pharmacokinetics of L-carnitine. Clin Pharmacokinet 2003;

42(11):941–67.

[79] Matsumoto Y, Amano I, Hirose S, Tsuruta Y, Hara S, Murata M, et al. Eﬀects of

L-carnitine supplementation on renal anemia in poor responders to erythropoietin. Blood
Purif 2001;19(1):24–32.

[80] Bellinghieri G, Santoro D, Calvani M, Mallamace A, Savica V. Carnitine and

hemodialysis. Am J Kidney Dis 2003;41(3 Suppl 1):S116–22.

[81] Evans A. Dialysis-related carnitine disorder and levocarnitine pharmacology. Am J

Kidney Dis 2003;41(4 Suppl 4):S13–26.

[82] Hurot JM, Cucherat M, Haugh M, Fouque D. Eﬀects of L-carnitine supplementation in

maintenance hemodialysis patients: a systematic review. J Am Soc Nephrol 2002;13(3):
708–14.

[83] Kittleson MD, Keene B, Pion PD, Loyer CG. Results of the multicenter spaniel trial

(MUST): taurine- and carnitine-responsive dilated cardiomyopathy in American cocker
spaniels with decreased plasma taurine concentration. J Vet Intern Med 1997;11(4):
204–11.

[84] Keene BW, Panciera DP, Atkins CE, Regitz V, Schmidt MJ, Shug AL. Myocardial L-

carnitine deﬁciency in a family of dogs with dilated cardiomyopathy. J Am Vet Med
Assoc 1991;198(4):647–50.

[85] Lofberg E, Essen P, McNurlan M, Wernerman J, Garlick P, Anderstam B, et al. Eﬀect of

hemodialysis on protein synthesis. Clin Nephrol 2000;54(4):284–94.

[86] Pion PD, Kittleson MD, Thomas WP, Delellis LA, Rogers QR. Response of cats with

dilated cardiomyopathy to taurine supplementation. J Am Vet Med Assoc 1992;201(2):
275–84.

[87] Suliman ME, Anderstam B, Bergstrom J. Evidence of taurine depletion and ac-

cumulation of cysteinesulﬁnic acid in chronic dialysis patients. Kidney Int 1996;50(5):
1713–7.

[88] Pion PD, Kittleson MD, Thomas WP, Skiles ML, Rogers QR. Clinical ﬁndings in cats

with dilated cardiomyopathy and relationship of ﬁndings to taurine deﬁciency. J Am Vet
Med Assoc 1992;201(2):267–74.

[89] Ross JR. Interventional nephrology. In: Johnson RJ, Fehally J, editors. Comprehensive

clinical nephrology. 2nd edition. Edinburgh: Mosby; 2003. p. 967–74.

[90] Beathard GA. Catheter thrombosis. Semin Dial 2001;14(6):441–5.
[91] Depner TA. Catheter performance. Semin Dial 2001;14(6):425–31.

966

J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

[92] Falk A, Mitty H, Guller J, Teodorescu V, Uribarri J, Vassalotti J. Thrombolysis of

clotted hemodialysis grafts with tissue-type plasminogen activator. J Vasc Interv Radiol
2001;12(3):305–11.

[93] Beathard GA. Mechanical versus pharmacomechanical thrombolysis for the treatment of

thrombosed dialysis access grafts. Kidney Int 1994;45(5):1401–6.

[94] Beathard GA. Gianturco self-expanding stent in the treatment of stenosis in dialysis

access grafts. Kidney Int 1993;43(4):872–7.

[95] Suhocki PV, Conlon PJ Jr, Knelson MH, Harland R, Schwab SJ. Silastic cuﬀed catheters

for hemodialysis vascular access: thrombolytic and mechanical correction of malfunction.
Am J Kidney Dis 1996;28(3):379–86.

[96] Lopot F, Nejedly B, Sulkova S, Blaha J. Comparison of diﬀerent techniques of

hemodialysis vascular access ﬂow evaluation. Int J Artif Organs 2003;26(12):1056–63.

[97] Kovalik EC, Schwab SJ. Treatment approaches for infected hemodialysis vascular

catheters. Curr Opin Nephrol Hypertens 2002;11(6):593–6.

[98] Marr KA, Sexton DJ, Conlon PJ, Corey GR, Schwab SJ, Kirkland KB. Catheter-related

bacteremia and outcome of attempted catheter salvage in patients undergoing hemodia-
lysis. Ann Intern Med 1997;127(4):275–80.

[99] Oliver MJ, Callery SM, Thorpe KE, Schwab SJ, Churchill DN. Risk of bacteremia from

temporary hemodialysis catheters by site of insertion and duration of use: a prospective
study. Kidney Int 2000;58(6):2543–5.

[100] Mokrzycki MH, Singhal A. Cost-eﬀectiveness of three strategies of managing tunnelled,

cuﬀed haemodialysis catheters in clinically mild or asymptomatic bacteraemias. Nephrol
Dial Transplant 2002;17(12):2196–203.

[101] Shaﬀer D. Catheter-related sepsis complicating long-term, tunnelled central venous

dialysis catheters: management by guidewire exchange. Am J Kidney Dis 1995;25(4):
593–6.

[102] Krishnasami Z, Carlton D, Bimbo L, Taylor ME, Balkovetz DF, Barker J, et al.

Management of hemodialysis catheter-related bacteremia with an adjunctive antibiotic
lock solution. Kidney Int 2002;61(3):1136–42.

[103] Droste JC, Jeraj HA, MacDonald A, Farrington K. Stability and in vitro eﬃcacy of

antibiotic-heparin lock solutions potentially useful for treatment of central venous
catheter-related sepsis. J Antimicrob Chemother 2003;51(4):849–55.

[104] Vercaigne LM, Sitar DS, Penner SB, Bernstein K, Wang GQ, Burczynski FJ. Antibiotic-

heparin lock: in vitro antibiotic stability combined with heparin in a central venous
catheter. Pharmacotherapy 2000;20(4):394–9.

[105] Adin CA, Cowgill LD. Treatment and outcome of dogs with leptospirosis: 36 cases

(1990–1998). J Am Vet Med Assoc 2000;216(3):371–5.

[106] Cowgill L, Francey T. Acute renal failure. In: Ettinger S, Feldman E, editors. Textbook

of veterinary internal medicine. 6th edition, in press.

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J.R. Fischer et al / Vet Clin Small Anim 34 (2004) 935–967

Update: management of calcium oxalate

uroliths in dogs and cats

Joseph W. Bartges, DVM, PhD*, Claudia Kirk, DVM,

PhD, India F. Lane, DVM, MS

Department of Small Animal Clinical Sciences, College of Veterinary Medicine, C247 Veterinary

Teaching Hospital, University of Tennessee, Knoxville, TN 39996–4544, USA

Incidence and types of uroliths

Urolithiasis is considered to be a common cause of lower urinary tract

disease in dogs and cats, and nephroliths seem to be diagnosed more
frequently than before

. Incidence rates for urolithiasis have not been

established for dogs and cats but are believed to be between 0.2% and 3.0%

[3,4]

. Many minerals may precipitate in the urinary tract, including, most

commonly, magnesium ammonium phosphate hexahydrate (struvite) and
calcium oxalate. In dogs and cats, the incidence of struvite urolithiasis is
decreasing, whereas the incidence of calcium oxalate urolithiasis is in-
creasing; it is the most common mineral found in feline uroliths (accounting
for 55% of feline uroliths submitted for quantitative analysis)

[4–6]

. In

addition, more than 90% of nephroliths and ureteroliths are composed of
calcium oxalate

Etiopathogenesis of uroliths

Overview

Urolith formation, dissolution, and prevention involve complex physical

processes. Major factors include (1) supersaturation of urine with calculo-
genic minerals resulting in crystal formation, (2) eﬀects of urinary inhibitors
of crystallization and urinary inhibitors of crystal aggregation and growth,
(3) urinary crystalloid complexors, (4) eﬀects of urinary promoters of crystal
aggregation and growth, and (5) eﬀects of noncrystalline matrix

[7,8]

. A

sequence of events leading to urolith formation is illustrated in

Fig. 1 [9]

* Corresponding author.
E-mail address:

jbartges@utk.edu

(J.W. Bartges).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.011

Vet Clin Small Anim

34 (2004) 969–987

Concept of urine saturation

An important driving force behind urolith formation is supersaturation of

urine with calculogenic substances

[7,9]

. This concept can be illustrated with

table salt (NaCl) and deionized water. When a salt is added to a solvent (eg,
NaCl in water), it dissolves in the solvent. The solution is said to be
undersaturated, because water contains such a low concentration of NaCl
that if more NaCl is added, it will dissolve. As more NaCl is added,
a concentration is reached beyond which further dissolution of NaCl is not
possible (

). At this point, water (solvent) is said to be saturated with

NaCl (salt). If additional NaCl is added to the water, saline (NaCl) crystals
will form unless the temperature or pH is changed. The concentration at
which saturation is reached and crystallization begins is called the thermo-
dynamic solubility product (see

;

). The thermodynamic solubility

product is a constant at which pure crystals of NaCl form in pure water.

Urine is a more complex solution than pure water, however. Urine

contains ions and proteins that interact or complex with calculogenic
minerals, such as calcium and oxalic acid, so as to allow them to remain in
solution. This explains why calcium and oxalic acid do not normally
precipitate in urine to form calcium oxalate crystals. Compared with water,
urine is normally supersaturated with respect to calcium and oxalic acid.

Saturation

Supersaturation

Nucleation

Crystal aggregation

or growth

Crystal retention

Stone formation

Calcium

Oxalate

Calcium
Oxalate

Fig. 1. Sequence of events leading to calcium oxalate urolith formation.

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

Energy is required to maintain this state of calcium and oxalic acid solubility;
therefore, urine must constantly ‘‘struggle’’ to maintain calcium and oxalic
acid in solution. Thus, urine is described as being metastable, implying
varying degrees of instability with respect to the potential for calcium oxalate
crystals to form (see

). In this metastable state, new calcium oxalate

crystals do not precipitate, but if they are already present, crystals can be
maintained and even grow in size. If the concentration of calcium or oxalic
acid is increased, a threshold is eventually reached at which urine cannot hold
more calcium and oxalic acid in solution. The urine concentration at which
this occurs is the thermodynamic formation product of calcium oxalate.
Above the thermodynamic formation product, urine is oversaturated and

1 teaspoon of salt dissolves completely (water is undersaturated with respect to salt)

3 teaspoons of salt dissolve completely, but any additional salt will result in precipitation (water is saturated with
respect to salt)

5 teaspoons of salt do not completely dissolve and results in precipitation. (water is supersaturated with respect
to salt)

Fig. 2. States of saturation illustrated by adding salt to water. (Top) One teaspoon of salt
dissolves completely (water is understaturated with respect to salt). (Middle) Three teaspoons of
salt dissolve completely, but any additional salt will result in precipitation (water is saturated
with respect to salt). (Bottom) Five teaspoons of salt do not completely dissolve, which results in
precipitation (water is supersaturated with respect to salt).

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

unstable with respect to calcium and oxalic acid. Thus, calcium oxalate
crystals spontaneously precipitate, grow in size, and aggregate.

Why is a conceptual understanding of these interrelated concepts of

clinical importance? Medical protocols for urolith dissolution result in
changing the state of oversaturation or metastability to a state of under-
saturation with calculogenic minerals. Medical treatment designed to
prevent urolith formation must create a state of undersaturation or at least
metastability as long as there is no mechanism for heterogeneous nucleation
present (see

Etiologic risk factors

Hypercalciuria

Excessive urinary excretion of calcium is a signiﬁcant contributor to

calcium oxalate formation in human beings and dogs. In human beings,

OVERSATURATION

(UNSTABLE SOLUTION)

Crystals spontaneously precipitate

(homogeneous nucleation)

Crystals aggregate and grow

Crystals do not dissolve

SUPERSATURATION

(METASTABLE SOLUTION)

Crystals do not spontaneously precipitate

Crystals precipitate on templates

(heterogeneous nucleation)

Crystals may aggregate

Inhibitors will impede or prevent

crystallization

Crystals do not dissolve

Thermodynamic formation product

UNDERSATURATION

(STABLE SOLUTION)

Crystals do not precipitate

Crystals do not aggregate or grow

Crystals dissolve

Thermodynamic solubility product

ENERGY FOR

DISSOLUTION

ENERGY FOR

PRECIPITATION

INCREASING

ACTIVITY
PRODUCT

INCREASING

IONIC

CONCENTRATION

Fig. 3. States of saturation.

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

however, 24-hour urinary excretion of calcium is normal in some calcium
oxalate urolith formers, and in some nonurolith-forming human beings, 24-
hour urinary excretion of calcium is elevated

. Hypercalciuria is thought

to be a risk factor but not necessarily the cause of calcium oxalate urolith
formation in human beings.

Calcium homeostasis is achieved through actions of parathyroid hor-

mone (PTH) and 1,25-dihydroxycholecalciferol (1,25-vitamin D) on bones,
intestines, and kidneys. When the serum ionized calcium concentration
decreases, PTH and 1,25-vitamin D activities increase, resulting in mobili-
zation of calcium from bone, increased absorption of calcium from intestine,
and increased reabsorption of calcium by renal tubules. Conversely, a high
serum ionized calcium concentration suppresses release of PTH and
production of 1,25-vitamin D, resulting in decreased bone mobilization,
decreased intestinal absorption of calcium, and increased urinary excretion
of calcium. Therefore, hypercalciuria can result from excessive intestinal
absorption of calcium (gastrointestinal [GI] hyperabsorption), impaired
renal reabsorption of calcium (renal leak), or excessive skeletal mobilization
of calcium (resorptive). In Miniature Schnauzers, GI hyperabsorption seems
to occur most commonly, although renal leak hypercalciuria has also been
observed

Consumption of diets supplemented with the urinary acidiﬁer ammonium

chloride by cats has been associated with increased urinary calcium excretion

. Additionally, consumption of diets containing high amounts of animal

protein by human beings results in metabolic acidosis and increased urinary
calcium excretion. Metabolic acidosis promotes hypercalciuria by promoting
bone turnover (release of calcium with buﬀers from bone), increased serum
ionized calcium concentration resulting in increased urinary calcium
excretion, and decreased renal tubular reabsorption of calcium. In dogs,
hypercalciuria resulting from ammonium chloride administration was de-
creased by bicarbonate administration

. In cats, magnesium supplemen-

tation as magnesium chloride was associated with increased urinary calcium
excretion and aciduria, whereas magnesium supplementation as magnesium
oxide was associated with alkaluria and a lesser degree of urinary calcium
excretion

[14]

. Magnesium may increase blood ionized calcium concentration

while suppressing PTH release.

Although excessive dietary intake of calcium may result in hypercalciuria,

studies in human beings refute this. Apparently, dietary calcium may bind to
dietary oxalic acid, resulting in calcium oxalate formation in the lumen of
the GI tract, thereby preventing absorption of calcium and oxalate.
Hypercalciuria may also occur with administration of loop diuretics,
glucocorticoids, urinary acidiﬁers, and vitamin D or C.

Hyperoxaluria

Oxalic acid is a metabolic end product of ascorbic acid and several amino

acids, such as glycine and serine, derived from dietary sources. Oxalic acid

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

forms soluble salts with sodium and potassium ions but a relatively
insoluble salt with calcium ions. Therefore, increased urinary excretion of
oxalic acid may promote calcium oxalate formation. Hyperoxaluria has
been observed in kittens consuming diets deﬁcient in vitamin B

(less than

1 mg/kg of diet)

. Hyperoxaluria has also been recognized in a group of

related cats with reduced quantities of hepatic D–glycerate dehydrogenase,
an enzyme involved in metabolism of oxalic acid precursors (primary
hyperoxaluria type II)

. Hyperoxaluria has also been associated with

defective peroxisomal alanine/glyoxylate aminotransferase activity (primary
hyperoxaluria type I) and intestinal disease in human beings (enteric
hyperoxaluria). These have not been identiﬁed in cats, and hyperoxaluria
has not been identiﬁed in dogs.

Altered inhibitors and promoters of calcium oxalate crystal formation

Urine is a complex solution containing many substances that may inhibit

or promote crystal formation and growth. Some inhibitors, such as citrate

, magnesium

[18]

, and pyrophosphate

, form soluble salts with

calcium or oxalic acid and thereby reduce the availability of calcium or
oxalic acid for precipitation. Other inhibitors are macromolecular proteins,
such as Tamm-Horsfall glycoprotein

and nephrocalcin

[21]

, which

interfere with the ability of calcium and oxalic acid to combine, thereby
minimizing crystal formation and growth. Other urinary substances may
promote calcium oxalate formation, including uric acid, which blocks
inhibitors of crystallization, or substances that serve as templates for
heterogeneous crystal nucleation, such as calcium phosphate crystals or
intraluminal suture material from a previous cystotomy. The role, if any, of
these crystallization inhibitors and promoters has not been evaluated in
dogs and cats.

Diagnosis of calcium oxalate uroliths

Historical information

Calcium oxalate uroliths tend to form in middle-aged to older animals;

however, Bichon Frisse dogs and Siamese cats seem to be predisposed to
forming calcium oxalate uroliths at a younger age (often at 3 to 4 years).
Historical information may include previous urinary tract disease or
underlying metabolic disease predisposing to urolith formation, or there
may be no preexisting clinical signs. When uroliths occur in the lower
urinary tract, clinical signs may include stranguria, hematuria, pollakiuria,
inappropriate urination, or urethral obstruction. Clinical signs associated
with uroliths that form in the kidneys or ureters may include polysystemic
illness (vomiting, depression, and anorexia) or abdominal pain, although
many upper urinary tract uroliths are not associated with clinical signs.
Calcium oxalate uroliths may form in association with other metabolic

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

diseases, such as diseases associated with hypercalcemia and with hyper-
adrenocorticism; clinical signs of the underlying disease may be most
obvious.

Physical examination

Physical examination is often normal unless urethral obstruction is

present. Urocystoliths may be palpated in approximately 20% of aﬀected
dogs and cats; it is more diﬃcult to palpate urocystoliths in dogs,
particularly large-breed dogs, than in cats. Dogs with hyperadrenocorticism
may show typical signs of the endocrinopathy.

Laboratory evaluation

Complete blood cell count and serum biochemical analysis

The complete blood cell count (CBC) and serum biochemical analysis are

usually normal. Hypercalcemia may be observed in approximately 4% of
dogs and 35% of cats with calcium oxalate uroliths

[4,5]

. In cats, total serum

calcium and ionized calcium are usually increased; however, the PTH
concentration is typically low. Azotemia, hyperkalemia, and metabolic
acidosis may be observed if urethral obstruction is present.

Urinalysis

Abnormalities may include hematuria, pyuria, bacteriuria, or crystalluria

(

. Urine pH is often acidic in animals with calcium oxalate

uroliths; however, if a urinary tract infection (UTI) with urease-producing
bacteria is present, alkaluria occurs in dogs and cats.

Fig. 4. Calcium oxalate dihydrate crystals in a dry mount urine sediment sample from a dog
with calcium oxalate uroliths. (

400 magniﬁcation).

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

Urine culture

Urine cultures often have no growth of bacteria but may be positive with

a secondary bacterial UTI. The most common organism causing bacterial
UTI is Escherichia coli.

Additional laboratory testing

Additional laboratory testing may be indicated in animals with predis-

posing metabolic diseases. Hyperadrenocorticism is a risk factor for de-
velopment of calcium oxalate uroliths; therefore, corticotropin stimulation
testing, low-dose dexamethasone suppression testing, high-dose dexametha-
sone suppression testing, or abdominal ultrasonography may be indicated.
Hypercalcemia is a risk factor for calcium oxalate formation; therefore,
measurement of blood ionized calcium concentration, PTH, and possibly
parathyroid-related protein concentration may be indicated. The most
common cause of hypercalcemia in dogs is neoplasia; however, calcium
oxalate uroliths do not usually form in these patients, most likely because
these animals show clinical signs of their neoplastic disease before uroliths
form. Primary hyperparathyroidism is an uncommon endocrine disorder but
has been associated with calcium oxalate urolith formation

[22,23]

. Hyper-

calcemia occurs in approximately one of three cats with calcium oxalate
uroliths; most often, the hypercalcemia is considered to be idiopathic. The
serum total calcium concentration and blood ionized calcium concentration
are increased; however, the PTH concentration is low

[24,25]

Imaging studies

Survey abdominal radiography is often suﬃcient for detection of calcium

oxalate uroliths because they are radiopaque (

)

.Use of double-

contrast cystography improves detection of urocystoliths. Small uroliths
may be retrieved during the procedure and submitted for quantitative
analysis. Excretory urography may be necessary to identify nephroliths and
ureteroliths and to determine if ureteral obstruction is present. Ultrasonog-
raphy may demonstrate the presence of uroliths, but it is diﬃcult to
determine the number and type of uroliths present (

). Additionally, the

ureters and urethra distal to the proximal part are often not visualized.

Tests of urine saturation

Preliminary evaluation in our laboratory has revealed that urinary

saturation with calcium oxalate is in the metastable range in clinically healthy
dogs and cats. Recall that urine in the metastable range is oversaturated with
calcium oxalate; however, crystals do not spontaneously form or precipitate
presumably because of the presence of crystallization inhibitors. In the few
cats with naturally occurring calcium oxalate uroliths that we have evaluated,
urinary saturation was above the thermodynamic formation product. These

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

tests may be useful in predicting urolith type, in evaluating risk of urolith
formation, and in monitoring management of uroliths.

Analysis of uroliths

Grossly, calcium oxalate dihydrate uroliths have a typical jagged surface

texture (

Fig. 7

); however, this is not true in all cases. Furthermore, calcium

oxalate monohydrate uroliths tend to have a smooth surface texture.
Quantitative analysis of uroliths voided during micturition or retrieved via
voiding urohydropropulsion, urinary catheterization, or cystotomy provides
the most information about the mineral composition of uroliths. When

Fig. 5. Lateral survey radiograph of calcium oxalate urocystoliths and a ureterolith in a 7-year-
old, spayed, female Miniature Schnauzer.

Fig. 6. Ultrasonographic image of a calcium oxalate urocystolith demonstrating acoustic
shadowing.

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

compared with quantitative analysis, qualitative analysis is correct in only
43% of the cases; therefore, quantitative analysis is recommended

Management of calcium oxalate uroliths

There is no medical protocol available for dissolution of calcium oxalate

uroliths; therefore, removal remains the treatment for uroliths that are
causing clinical problems

[4,5]

Urocystoliths that cause repeated urethral obstruction or clinical signs or

are associated with bacterial UTI should be treated. Likewise, nephroliths
that cause ureteral obstruction; continue to grow in size or number and
damage renal tissue; or cause recurrent bacterial UTI, severe hematuria, or
pain should be treated. Many animals may live with uroliths for months or
years with minimal or no clinical signs; therefore, the decision to treat uroliths
should be discussed with the owner. Risks and beneﬁts of medical (

versus surgical (

) therapy of uroliths should be considered. Urocysto-

liths that are smaller than the smallest luminal diameter of the urethra may be
retrieved using voiding urohydropropulsion

or a catheter-assisted

retrieval technique

. Because calcium oxalate uroliths are recurrent,

removal of uroliths should not be the end point of therapy. Appropriate
preventative measures and follow-up evaluations should be instituted.

Fig. 7. Gross appearance of calcium oxalate dihydrate urocystoliths from a 10-year-old, male,
castrated cat (left) and calcium oxalate monohydrate urocystoliths from a 9-year-old, male,
castrated Dachshund (right).

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Retrograde urohydropropulsion

Uroliths that cause urethral obstruction should be retropulsed into the

urinary bladder

. This can be accomplished as described. Make a dilute

sterile lubricant solution by mixing one part of sterile lubricant with
approximately two parts of sterile water or crystalloid solution. This is done
by placing sterile lubricant (5 mL) in a 35-mL syringe, attaching a three-way
stopcock, and placing sterile ﬂuid (10–15 mL) in a second 35-mL syringe.
Attach the syringes to the three-way stopcock, squirt the sterile solution into
the lubricant, and inject the sterile lubricant-ﬂuid solution back into the
empty syringe. Repeat several times to mix the lubricant and ﬂuid. Sedate or
anesthetize the dog or cat. Insert a red rubber catheter (5.0–8.0 French) in
dogs or a polypropylene catheter (3.5 French) in cats to the site of
obstruction. Infuse sterile lubricant-ﬂuid solution to lubricate the urethra.
Sometimes, this is all that is needed to retropulse urethroliths into the
bladder. If the uroliths do not move, attach a syringe ﬁlled with sterile ﬂuid to
the urethral catheter. The urethra is occluded proximally by trapping the
urethra against the ﬂoor of the pelvis per the rectum. At the same time,
occlude the distal penile urethra and infuse sterile ﬂuid under pressure. When
the urethra is distended, release occlusion of the pelvic urethra. If uroliths are
retropulsed into the bladder, there should be a ‘‘popping’’ sensation and the

Box 1. Overview of medical management of uroliths

1. Institute appropriate medical or surgical therapy. Consider

surgical correction if uroliths are obstructing urine flow or if
correctable abnormalities predisposing to recurrent uroliths
are identified.

2. Eradicate or control urinary tract infection (UTI).
3. Initiate therapy with appropriate diet with or without

appropriate pharmacologic therapy.

4. Devise a protocol to monitor the efficacy of therapy.

A. Try to avoid diagnostic follow-up studies that require

urinary catheterization. If they are required, give
appropriate pericatheterization antimicrobial agents to
prevent iatrogenic UTI.

B. Evaluate serial urinalyses. Urine pH, urine specific

gravity (USPG), and microscopic examination
of sediment for crystals and bacteria are important.
Remember crystals formed in urine stored at room
or refrigeration temperature may represent in vitro
artifacts.

C. Perform serial radiography to evaluate urolith

location(s), number, size, density, and shape.

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

Box 2. Overview of general principles and considerations
for surgical treatment of uroliths

Preoperative considerations

1. Obtain all blood and urine samples before administration of

diagnostic (radiopaque contrast media) or therapeutic (eg,
antibiotics, fluids) agents. Quantitatively evaluate renal
function.

2. Always radiograph the entire urinary tract to determine the

location and number of uroliths. Contrast radiography or
ultrasonography should be considered to evaluate the
patency of the excretory pathway. Thoroughly evaluate the
entire urinary tract for correctable anatomic abnormalities
that may have initiated urinary infection and subsequent
struvite urolithiasis.

Operative considerations

1. Preserve renal organ function. Avoid use of mattress sutures

to repair nephrotomy incisions because they cause
additional irreversible loss of renal function because of
infarction.

2. Remove nephroliths before removing cystoliths in patients

with multiple uroliths. If ureteroliths are present or if small or
fractured nephroliths subsequently pass into the ureters,
they may be flushed into the bladder.

3. Obtain urine samples for routine analysis and microbial

culture if they could not be obtained before surgery. Obtain
biopsy samples of the urinary tract at the time of
nephrotomy, pyelotomy, cystotomy, or urethrotomy.
Evaluation of biopsy samples may be of diagnostic and
prognostic significance.

4. Make a special effort to remove all uroliths. Thoroughly flush

the affected lumen of the urinary tract with a sterile isotonic
solution to remove small uroliths. Bacteria harbored inside
struvite uroliths allow urinary tract infections to persist and
predispose to recurrence of struvite uroliths.

5. When possible, do not allow suture material to penetrate the

lumen of the urinary tract. Suture material may serve as
a nidus for urolith formation by lowering the formation
product. Nonabsorbable and multifilament sutures are more
calculogenic than absorbable or monofilament sutures.

6. Save all uroliths for mineral analysis, possible culture, and

possible microscopic examination. Culture of uroliths may
help to detect a bacterial organism in patients receiving

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

person occluding the urethra per the rectum can often feel the uroliths move
cranially into the bladder. Most uroliths can be moved successfully with this
procedure. Urate, cystine, and most struvite uroliths move easily because of
their smooth texture; calcium oxalate uroliths are less mobile because of their
irregular surface texture. This procedure will not be successful if uroliths are
embedded in the urethral mucosa or if there is a stricture proximal to the
uroliths’ location in the urethra.

Voiding urohydropropulsion

Voiding urohydropropulsion may be used to retrieve uroliths that have

a diameter smaller than the smallest diameter of the urethral lumen.
Typically, uroliths smaller than approximately 5 mm in female cats, 1 mm
in male cats, 10 to 15 mm in female dogs, and 1 to 3 mm in male dogs can be
retrieved. For larger uroliths, a cystotomy must be performed. Voiding
urohydropropulsion is performed as follows

. The dog or cat is usually

anesthetized or heavily sedated. The urinary bladder should be palpably
distended. If it is not, insert a urinary catheter and distend the bladder with
a sterile crystalloid solution. The dog or cat is then lifted and supported under
the axillary region so that the vertebral column is perpendicular to the

antimicrobial therapy before diagnostic urine culture and
antimicrobial susceptibility tests are performed.

Postoperative considerations

1. Avoid use of indwelling catheters because they are

a common cause of iatrogenic urinary tract infections. If one
must use an indwelling catheter, a closed system should be
used when possible because it minimizes retrograde
migration of pathogens through the catheter lumen.

2. If multiple uroliths are present, evaluate the urinary tract

radiographically after surgery. Immediate detection of
uroliths that were inadvertently allowed to remain in the
urinary tract is of prognostic significance. It may be
erroneously assumed that the patient is highly predisposed
to recurrent urolithiasis if residual uroliths are first detected
on radiographs taken several weeks after surgery.
Appropriate medical or surgical therapy should be
formulated to manage residual uroliths.

3. Therapy should be designed to promote postoperative

diuresis in patients undergoing nephrolithotomy. Increased
urine flow minimizes the formation of blood clots in the renal
pelves, which have the potential to obstruct urine outflow or
to mineralize.

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ground. The urinary bladder is then palpated and gently agitated to dislodge
any uroliths adhering to the bladder mucosa and to position uroliths in the
urinary bladder trigone. The urinary catheter is removed, and a collection
container is positioned under the vulva or tip of the penis. Uroliths are
expelled by gently applying pressure to the urinary bladder so that the
increase in intraluminal pressure initiates a micturition reﬂex. Uroliths are
not squeezed out in this procedure; they are expelled by micturition. Repeat
the procedure if the number of uroliths voided is less than that previously
detected by radiography. If uroliths detected by radiography were too
numerous to count, repeat voiding urohydropropulsion until uroliths are
no longer observed in the expelled ﬂuid. If the number of uroliths retrieved
equals the number observed by radiography, recover the patient. If there is
a question concerning whether all the uroliths were retrieved, sequential
survey abdominal radiography can be used to monitor progress. Animals
should be treated for 3 to 5 days with antibiotics because of the possible
introduction of bacteria by catheterization. We recommend repeating
urinalysis and a urine culture 5 to 10 days after discontinuation of antibiotic
treatment to ensure eradication of infection.

Nephroliths and ureteroliths

Nephroliths and ureteroliths are commonly composed of calcium oxalate.

Because calcium oxalate uroliths cannot be dissolved using medical proto-
cols, surgical removal or lithotripsy is the only option if the urolith must be
removed. Many times, nephroliths or renal mineral densities are observed
incidentally on survey abdominal radiographs; however, in animals that are
hematuric or have renal failure or nonspeciﬁc abdominal pain, nephroliths or
ureteroliths may be the cause. The decision to remove a nephrolith or
ureterolith should be considered carefully because of the diﬃculty associated
with ureteral surgery in cats and the long-term damage to a kidney induced by
nephrotomy. Upper urinary tract uroliths should be removed if they are
causing obstruction resulting in diminished renal function; if they are
associated with severe hematuria, pain, or persistent bacterial infection; or
if they are increasing in size and damaging renal tissue despite appropriate
medical management (see section on prevention). If none of these conditions
is present, a reasonable alternative approach includes preventative measures
to minimize an increase in the size or number of uroliths. Monitoring may be
accomplished by performing survey abdominal radiography every 3 to
6 months. If uroliths increase in size or number or cause pain, hematuria,
infection, or obstruction, direct treatment (surgery or lithotripsy) should be
considered to prevent loss of that kidney.

Surgical management

Problematic ureteroliths and nephroliths must be removed by surgery or

lithotripsy if renal function is to be protected or preserved. A modiﬁed

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nephrotomy or pyelolithotomy is used for nephroliths or proximal uretero-
liths. Nephrectomy is performed with severely hydronephrotic, infected, or
nonfunctional kidneys. Resection and reimplantation of ureters are eﬀective
for surgical management of distal ureteroliths. Ureterotomy has been
successful for removal of obstructive ureteroliths in some dogs and cats,
but this approach requires a more experienced surgical and postoperative
team. Microscopic surgical techniques can be used to remove obstructive
ureteroliths with good long-term outcomes in cats

. If a distal ureterolith is

moveable, it may be advanced into the bladder and removed by cystotomy.

Lithotripsy

Lithotripsy refers to ‘‘breaking of stones’’ and oﬀers a nonsurgical

modality for the management of uroliths. Indications for lithotripsy are
similar to those for surgical intervention for upper tract uroliths. Lithotripsy
may be preferred for nephroliths in solitary kidneys, for ureteroliths, or for
animals that are poor surgical candidates. Overall, lithotripsy is considered
less damaging to renal tissue and function than nephrotomy and oﬀers a good
alternative to open surgery

. Lithotripsy treatment of ureteroliths is a bit

more diﬃcult because of more diﬃcult positioning and the limited movement
of impacted ureteroliths within the treatment ﬁeld. Successful fragmentation
of ureteroliths may require multiple treatments. Ureteroscopic techniques
now are favored by some for the management of distal ureteral uroliths in
human beings and may be adaptable to large dogs in the future. Although
much of the original extracorporeal shock wave lithotripsy (ESWL) research
included animal models, including dogs, experience with treatment of clinical
uroliths in dogs and cats has been limited

[30–32]

. Additional discussion of

the procedures and outcomes of lithotripsy can be found in another article in
this issue.

Combination therapy of uroliths

Diﬀerent modalities of urolith management may be used in combination.

Occasionally, uroliths are missed at the time of cystotomy. Calcium oxalate
uroliths are incompletely removed at the time of cystotomy in 15% to 20%
of dogs and cats

. If urocystoliths remain after cystotomy and are small

enough in diameter to pass, they may be retrieved using voiding urohy-
dropropulsion. If uroliths are too large to recover by nonsurgical means,
they must be removed surgically or steps to minimize the growth of the
uroliths must be undertaken.

Prevention of uroliths

Because the causes of calcium oxalate urolith formation are not completely

known, no treatment has been shown to be completely eﬀective (

Box 3

;

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J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

Box 3. Summary of recommendations for medical treatment of
calcium oxalate uroliths

1. Perform appropriate diagnostic studies including complete

urinalysis, quantitative urine culture, serum biochemical profile,
and diagnostic radiography or ultrasonography. Determine
precise location, size, and number of uroliths, and patency of
excretory pathway.

2. If available, determine the mineral composition of uroliths. If

unavailable, ‘‘guesstimate’’ their composition by evaluation of
appropriate clinical data.

3. Determine urine concentrations of appropriate metabolites (if

possible), especially calcium, oxalate, magnesium, uric acid, and
citrate.

4. Consider immediate surgical correction if uroliths thought to be

composed of calcium oxalate are causing intolerable clinical
signs. Uroliths may also be removed by catheter-assisted
retrieval or by voiding urohydropropulsion.

5. If necessary, eradicate or control secondary urinary tract

infections with appropriate antimicrobial agents.

6. Hypercalcemic hypercalciuric patients

A. Approximately 35% of cats and 4% of dogs with calcium

oxalate uroliths are hypercalcemic.

B. Have a high index of suspicion of primary

hyperparathyroidism. If confirmed, surgically correct
abnormality of parathyroid glands. This is a rare cause
of calcium oxalate uroliths.

C. If uroliths are symptomatic, consider surgical removal.
D. Avoid hydrochlorothiazide because it may aggravate

hypercalcemia.

E. Induce polyuria (but avoid excessive dietary sodium

supplements).

High-fiber diets seem to decrease the degree of
hypercalcemia and should be used with potassium citrate
(initial dose: 75 mg/kg administered orally [PO] every
12 hours; adjust dose to induce a urine pH of 7.0 to 7.5).

7. Normocalcemic patients with active calcium oxalate urolithiasis

A. Feed a protein-restricted and alkalinizing diet.
B. Consider oral administration of potassium citrate (see

above).

C. Consider change to a diet that does not contain excessive

oxalate, sodium, or protein.

D. Avoid dietary or therapeutic supplements of ascorbic acid

and vitamin D.

E. Consider hydrochlorothiazide (2–4 mg/kg PO every 12 hours)

or vitamin B

(2 mg/kg PO every 24 hours).

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). Calcium oxalate uroliths are recurrent; thus, preventative measures

are warranted to prevent formation of uroliths. Production of dilute urine is
probably beneﬁcial in dogs and cats with calcium oxalate urolithiasis.

In dogs, feeding a protein- and sodium-restricted alkalinizing diet has

been shown to delay recurrence

. If a neutral to slightly alkaline urine pH

is not accomplished by diet alone, potassium citrate may be administered
(initial dose: 75 mg/kg administered orally [PO] every 12 hours; adjust to
induce a urine pH of 7.0 to 7.5). Because the protein- and sodium-restricted,
alkalinizing diet is also high in fat, some dogs cannot tolerate the diet. In
these dogs, feeding a low-fat higher ﬁber diet with supplemental potassium
citrate seems to be eﬀective.

In cats, several diets are available that are formulated to reduce calcium

and oxalic acid concentrations in urine, promote high concentration and
activity of inhibitors of calcium oxalate crystal growth and aggregation in
urine, and maintain dilute urine. Consumption of these diets by healthy cats
results in production of urine that is undersaturated with calcium oxalate

[33,34]

. In one study of cats with naturally occurring calcium oxalate

uroliths, consumption of one ‘‘oxalate preventative diet’’ resulted in
a decrease of urine saturation from the oversaturated state to a metastable
state; cats did not reform uroliths

. In cats with hypercalcemia and

calcium oxalate uroliths, prevention seems to be more successful when
feeding a higher ﬁber-containing diet and administering potassium citrate
(initial dose: 75 mg/kg PO every 12 hours; adjust to induce a urine pH of 7.0
to 7.5)

. Other treatments that have been proposed include vitamin B

(2 mg/kg PO every 24 hours) and hydrochlorothiazide (2–4 mg/kg PO every
12 hours).

Calcium oxalate uroliths are recurrent. Serial monitoring of a dog or cat

with a history of calcium oxalate urolithiasis should be part of the
preventative protocol. Periodically, a complete urinalysis should be per-
formed to monitor urine speciﬁc gravity, pH, and the presence of calcium

Table 1
Expected changes with therapy of calcium oxalate uroliths

Factor

Pretherapy

Prevention

Polyuria

Variable

Pollakiuria

0 to 4

Hematuria

0 to 4

USPG

Variable

1.004–1.025

Urine pH

\ 7.0

>7.0

Urine inﬂammation

0 to 4

Calcium oxalate crystals

0 to 4

Bacteriuria

0 to 4

Culture

Positive/negative

Negative

BUN (mg/dl)

>15

10–30

Urolith size and number

Small to large

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oxalate crystalluria. Ideally, urine should be dilute, urine pH should be in
the neutral to alkaline range, and calcium oxalate crystalluria should not be
present. Survey abdominal radiography should be performed approximately
every 6 months to evaluate for recurrence. If calcium oxalate urocystoliths
are detected while small in size, they may be retrieved nonsurgically, and
adjustment to the preventative protocol can be made.

References

[1] Ross SJ, Osborne CA, Lulich JP, et al. Canine and feline nephrolithiasis. Epidemiology,

detection, and management. Vet Clin North Am Small Anim Pract 1999;29:231–50.

[2] Lekchardensuk C, Osborne CA, Lulich JP, et al. Increased frequency of calcium oxalate

uroliths in the upper urinary tract of cats: 1981 to 1999. In: Managing urolithiasis in cats:
recent updates and practice guidelines. Topeka (KS): Hill’s Pet Nutrition; 2003. p. 24–5.

[3] Osborne CA, Bartges JW, Lulich JP, et al. Canine urolithiasis. In: Hand MS, Thatcher

CD, Remillard RL, Roudebush P, editors. Small animal clinical nutrition. 4th edition.
Marceline, MO: Wadsworth Publishing Co.; 2000. p. 605–88.

[4] Osborne CA, Kruger JM, Lulich JP, et al. Feline lower urinary tract diseases. In: Ettinger

SJ, Feldman EC, editors. Textbook of veterinary internal medicine. 5th edition.
Philadelphia: WB Saunders; 1999. p. 1710–46.

[5] Lulich JP, Osborne CA, Bartges JW, et al. Canine lower urinary tract disorders. In:

Ettinger SJ, Feldman EC, editors. Textbook of veterinary internal medicine. 5th ed.
Philadelphia: WB Saunders; 1999. p. 1747–83.

[6] Osborne CA, Lulich JP, Albansan H, et al. Mineral composition of feline uroliths and

urethral plugs: current status. In: Managing urolithiasis in cats: recent updates and practice
guidelines. Topeka (KS): Hill’s Pet Nutrition; 2003. p. 26–8.

[7] Coe FL, Parks JH, Asplin JR. The pathogenesis and treatment of kidney stones. N Engl J

Med 1992;327:1141–52.

[8] Brown C, Purich D. Physical-chemical processes in kidney stone formation. In: Coe F,

Favus M, editors. Disorders of bone and mineral metabolism. New York: Raven Press;
1992. p. 613–24.

[9] Bartges JW, Osborne CA, Lulich JP, et al. Methods for evaluating treatment of uroliths.

Vet Clin North Am Small Anim Pract 1999;29:45–57.

[10] Wilson DM. Clinical and laboratory evaluation of renal stone patients. Endocrinol Metab

Clin North Am 1990;19:773–803.

[11] Lulich JP, Osborne CA, Nagode LA, et al. Evaluation of urine and serum metabolites in

miniature schnauzers with calcium oxalate urolithiasis. Am J Vet Res 1991;52:1583–90.

[12] Ching SV, Fettman MJ, Hamar DW, et al. The eﬀect of chronic dietary acidiﬁcation using

ammonium chloride on acid-base and mineral metabolism in the adult cat. J Nutr 1989;
119:902–15.

[13] Sutton RA, Wong NL, Dirks J. Eﬀects of metabolic acidosis and alkalosis on sodium and

calcium transport in the dog kidney. Kid Int 1979;15:520–33.

[14] Buﬃngton CA, Rogers QR, Morris JG. Eﬀect of diet on struvite activity product in feline

urine. Am J Vet Res 1990;51:2025–30.

[15] Bai SC, Sampson DA, Morris JG, et al. Vitamin B6 requirement of growing kittens. J Nutr

1989;119:1020–7.

[16] McKerrell RE, Blakemore WF, Heath MF, et al. Primary hyperoxaluria (L-glyceric

aciduria) in the cat: a newly recognised inherited disease. Vet Rec 1989;125:31–4.

[17] Lieske JC, Leonard R, Toback FG. Adhesion of calcium-oxalate monohydrate crystals to

renal epithelial-cells is inhibited by speciﬁc anions. Am J Physiol Renal Fluid Electrolyte
Physiol 1995;37:F604–12.

986

J.W. Bartges et al / Vet Clin Small Anim 34 (2004) 969–987

[18] Tur JA, Prieto R, Grases F. An animal-model to study the eﬀects of diet on risk-factors of

calcium stone formation. Scand J Urol Nephrol 1991;25:311–4.

[19] Russell RG, Bisaz S, Fleisch H. The inﬂuence of orthophosphate on the renal handling of

inorganic pyrophosphate in man and dog. Clin Sci Mol Med 1976;51:435–43.

[20] Gokhale JA, Glenton PA, Khan SR. Characterization of Tamm-Horsfall protein in a rat

nephrolithiasis model. J Urol 2001;166:1492–7.

[21] Nakagawa Y, Ahmed M, Hall SL, et al. Isolation from human calcium oxalate renal stones

of nephrocalcin, a glycoprotein inhibitor of calcium oxalate crystal growth. Evidence that
nephrocalcin from patients with calcium oxalate nephrolithiasis is deﬁcient in gamma-
carboxyglutamic acid. J Clin Invest 1987;79:1782–7.

[22] Marquez GA, Klausner JS, Osborne CA. Calcium oxalate urolithiasis in a cat with

a functional parathyroid adenocarcinoma. J Am Vet Med Assoc 1995;206:817–9.

[23] Klausner JS, O’Leary TP, Osborne CA. Calcium urolithiasis in two dogs with parathyroid

adenomas. J Am Vet Med Assoc 1987;191:1423–6.

[24] Midkiﬀ AM, Chew DJ, Randolph JF, et al. Idiopathic hypercalcemia in cats. J Vet Intern

Med 2000;14:619–26.

[25] Savary KC, Price GS, Vaden SL. Hypercalcemia in cats: a retrospective study of 71 cases

(1991–1997). J Vet Intern Med 2000;14:184–9.

[26] Lulich JP, Osborne CA, Sanderson SL, et al. Voiding urohydropropulsion: lessons from

5 years of experience. Vet Clin North Am Small Anim Pract 1999;29:283–92.

[27] Lulich JP, Osborne CA. Catheter-assisted retrieval of urocystoliths from dogs and cats.

J Am Vet Med Assoc 1992;201:111–3.

[28] Osborne CA, Lulich JP, Polzin DJ, et al. Medical dissolution and prevention of canine

struvite urolithiasis. Twenty years of experience. Vet Clin North Am Small Anim Pract
1999;29:73–111.

[29] Kyles AE, Stone EA, Gookin J, et al. Diagnosis and surgical management of obstructive

ureteral calculi in cats: 11 cases (1993–1996). J Am Vet Med Assoc 1998;213:1150–6.

[30] Adams LG, Senior DF. Electrohydraulic and extracorporeal shock-wave lithotripsy. Vet

Clin North Am Small Anim Pract 1999;29:293–302.

[31] Bailey G, Burk RL. Dry extracorporeal shock wave lithotripsy for treatment of

ureterolithiasis and nephrolithiasis in a dog. J Am Vet Med Assoc 1995;207:592–5.

[32] Block G, Adams LG, Widmer WR, et al. Use of extracorporeal shock wave lithotripsy for

treatment of nephrolithiasis and ureterolithiasis in ﬁve dogs. J Am Vet Med Assoc 1996;
208:531–6.

[33] Smith BHE, Moodie SJ, Wensley S, et al. Diﬀerences in urinary pH and relative

supersaturation values between senior and young adult cats. Presented at the 15th
American College of Veterinary Internal Medicine Forum. Orlando, FL, 1997.

[34] Smith BH, Stevenson AE, Markwell PJ. Urinary relative supersaturations of calcium

oxalate and struvite in cats are inﬂuenced by diet. J Nutr 1998;128(Suppl):2763S–4S.

[35] Lulich JP, Osborne CA, Lekcharoensuk C, et al. Eﬀects of diets on composition of urine of

cats with calcium oxalate urolithiasis. J Am Anim Hosp Assoc 2004;40:185–91.

[36] McClain HM, Barsanti JA, Bartges JW. Hypercalcemia and calcium oxalate urolithiasis in

cats: a report of ﬁve cases. J Am Anim Hosp Assoc 1999;35:297–301.

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Management of ureteral obstruction

Elizabeth M. Hardie, DVM, PhD

Andrew E. Kyles, BVMS, PhD

Department of Clinical Sciences, College of Veterinary Medicine, North Carolina

State University, 4700 Hillsborough Street, Raleigh, NC 27606, USA

Department of Surgical and Radiological Sciences, School of Veterinary Medicine,

1410 Tupper Hall, University of California at Davis, Davis, California 95616, USA

Ureteral obstruction leads to restriction of urine ﬂow, which may result in

uremic crisis, changes in the structure of the kidney and ureter, and loss of
renal function. Relief of obstruction in a timely fashion can preserve the
structure and function of the kidneys. Small animal practitioners need to be
familiar with the clinical signs related to obstruction, the consequences of
obstruction, and the techniques used to restore patency to the ureter.

Anatomy

The ureter is a retroperitoneal structure that joins the renal pelvis and the

bladder, serving as a conduit for urine

. The ureter is lined with

transitional cell epithelium surrounded by a connective tissue layer called
the lamina propria. Together, these two layers make up the mucosa, which
lies in folds (

). The lumen is normally collapsed, opening only when

the bolus of urine passes through. Surrounding the mucosa are several layers
of smooth muscle that function as a syncytium

. The normal pacemaker

for this muscle lies within the kidney. When the pacemaker depolarizes, the
muscle contracts in peristaltic waves, propelling urine from cranial to
caudal. Surrounding the muscle is a loose layer of adventitia

. The

adventitia contains fat, the ureteral vessels, and lymphatics. The cranial
ureteral artery comes from the renal artery, whereas the caudal ureteral
artery comes from the prostatic/vaginal artery. There may be contributions
from the testicular/ovarian vessels, the iliac vessels, and the vesicular vessels

[1,4]

. The ureter is well innervated with sympathetic, parasympathetic, and

* Corresponding author.
E-mail address:

lizette_hardie@ncsu.edu

(E.M. Hardie).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.008

Vet Clin Small Anim

34 (2004) 989–1010

sensory nerves, but nerve function is not necessary for the peristaltic
function of the ureter

The renal end of the ureter (ureteropelvic junction) is covered by renal

parenchyma in the dog and cat

[1,4,5]

. Once the ureter leaves the kidney, it

passes dorsal to the testicular/ovarian vessels. The right ureter may course
dorsal or lateral to the vena cava, whereas the left ureter is usually lateral to
the aorta. As the ureter continues caudally, it passes ventral to the deep
circumﬂex iliac and external iliac vessels. In male animals, it crosses dorsal
to the ductus deferens, whereas in female animals, it courses in the dorsal
aspect of the broad ligament. The ureter then enters the lateral ligament of
the bladder. The distal end of the ureter curves before entering the bladder,
resulting in a J-shaped hook conﬁguration

. The ureter enters the bladder

wall on the dorsocaudolateral aspect. The mucosa and lamina propria
continue through the bladder wall, but the muscle layer is replaced by
attachments from the detrusor muscle

. The ureter runs in the submucosa

before opening into the bladder lumen. When the bladder is full, this region
of the ureter is compressed, preventing urine from being forced into the
ureter from the bladder.

In human beings, three sites of potential narrowing and obstruction have

been identiﬁed: the ureteropelvic junction, the site at which the ureter
crosses the iliac vessels, and the ureterovesical junction

. Similar sites have

not been identiﬁed for the dog and cat, but it seems likely that they exist.
The normal diameter of the canine ureter, as measured using helical CT,
ranges from 1.3 to 2.7 mm

. Spheres with a diameter of 2.3 mm can be

passed down the canine ureter, whereas spheres with a diameter greater than

Fig. 1. Section of normal ureter from an adult cat. The section is from the midureter region.
Note the folded mucosa surrounded by smooth muscle and the abundant periureteral adipose
tissue. Hematoxylin-eosin section with bar = 100 lm (0.1 mm). (Courtesy of Dr. Stephen
Griﬀey, Davis, CA.)

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or equal to 2.8 mm lodge in the lumen

. The normal outside diameter of

the feline ureter is 1 mm, whereas the lumen is 0.4 mm

[4,8]

. Suture with

a diameter of 0.3 to 0.4 mm (US Pharmacopeia size 2-0 to 0) can usually be
passed down the lumen of a feline ureter. The lumen of the ureter can dilate
up to 17 times normal in response to diuresis

Physiology

The physiologic response to ureteral obstruction is extremely complex and

depends on the species, age of the animal, degree of obstruction, length of time
the obstruction exists, and whether or not the obstruction is unilateral or
bilateral

. After complete unilateral ureteral obstruction, renal blood ﬂow

and ureteral pressure increase for 1 to 1.5 hours on the aﬀected side. The blood
ﬂow then begins to decline, whereas ureteral pressure continues to increase.
After 5 hours, renal blood ﬂow decreases further and ureteral pressure
decreases. Ureteral pressure is near normal by 24 hours after obstruction.
Renal blood ﬂow continues to decrease, and by 2 weeks after obstruction,
renal blood ﬂow is 20% of normal in conscious dogs

. In dogs, the

glomerular ﬁltration rate (GFR) increases in the aﬀected kidney initially but
then decreases

. The GFR in the contralateral kidney increases.

After relief of complete unilateral ureteral obstruction, return of the GFR

ranges from nearly normal after 4 days of obstruction to 46% of normal after 2
weeks

. Return of concentrating ability can occur if obstruction is less than

1 week but is permanently impaired after 4 weeks of obstruction

Interestingly, destruction of the obstructed kidney occurs more quickly and
return of function occurs more slowly if the contralateral kidney is present and
functioning

. In contrast to the rapid irreversibility of renal function in

complete obstruction, partial obstruction results in less severe destruction and
more return of function after relief of obstruction. In one model using dogs,
total GFR was normal after 4 weeks of partial obstruction

Congenital ureteral obstruction can lead to renal dysplasia, whereas

gradual renal atrophy and destruction occur with long-standing obstruction
in adults. By 18 months in dogs, the kidney is replaced by a thin-walled sac
of ﬂuid

. In the ureter, obstruction leads to hydroureter and thickening of

the smooth muscle layer. Thickening is caused by muscular hypertrophy
rather than by hyperplasia, and the muscle is gradually replaced by ﬁbrous
tissue

. More than 90% of the muscle layer is ﬁbrous tissue by 42 days

after obstruction in the rat, but the exact time this takes in dogs and cats is
not known.

Causes of obstruction

Mechanical obstruction of the ureter can result from an intraluminal

obstruction, mural lesion, or extraluminal compression. Common causes of

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ureteral obstruction in dogs and cats include ureteral calculi, neoplasia,
trauma, inﬂammation, ﬁbrosis, congenital stenosis, acquired stricture,
foreign bodies, and blood clots

Congenital ureteral obstruction is a rarely reported condition in the dog.

Bilateral ureteropelvic obstruction

has been reported in a puppy,

leading to the death of the dog. It is likely that similar conditions probably
lead to unexplained neonatal deaths, but unless a full necropsy is performed,
the cause may be missed. Unilateral segmental ureteral aplasia has been
described associated with ectopic ureter

Calculi are the most commonly reported cause of intraluminal ureteral

obstruction in the dog but are still rare (51 of 13,548 urinary calculi
examined by the Urinary Stone Analysis Laboratory at the University of
California at Davis between 1981 and 1995)

[14]

. Struvite and calcium

oxalate calculi have been removed from the ureter

[15,16]

. Debris from

pyelonephritis has also been reported to occlude the ureter

Mural causes of obstruction in the dog include ureterocele

[18,19]

ﬁbrosis

, ﬁbroepithelial polyps

, proliferative ureteritis

ureteral neoplasm, and metastasis

. Fibrosis has been associated with

radiation therapy. Fibroepithelial polyps and proliferative ureteritis are
benign polypoid lesions. Histologically, they contain a layer of normal
epithelial cells covering a prominent ﬁbrovascular stroma with lymphoplas-
macytic inﬂammation. It is unknown whether these lesions are a form of
chronic inﬂammation or a benign tumor. Reported ureteral neoplasms
include ﬁbropapilloma, leiomyoma, leiomyosarcoma, and primary ureteral
transitional cell carcinoma. Metastasis from other sites of transitional cell
carcinoma can also occur.

Extramural causes of ureteral obstruction in the dog include pelvic

masses, bladder neoplasms, prostatic neoplasms, and ligatures

[5,13]

. Any

mass that compresses or envelops the ureter can result in obstruction. Poor
placement of ligatures associated with ovariohysterectomy can result in
obstruction caused by entrapment of the ureter in the ligature or in scar
tissue associated with the ligature. Most often, these ligatures result in
complete obstruction; however, rarely, partial obstruction caused by scar
tissue can result in a more chronic course of disease

[24,25]

. One dog was

presented 7 years after ovariohysterectomy for renal failure with associated
bilateral hydroureter and hydronephrosis

The most common cause of ureteral obstruction in cats is ureteral calculi

[26–31]

. Ureteral calculi are being diagnosed with increasing frequency in

cats

. A search of the computerized medical record system at the

University of California at Davis revealed that the ﬁrst cat was diagnosed
with ureteral calculi in 1990, and there has been a gradual progressive
increase in the prevalence of ureteral calculi, with 23 cats being diagnosed in
2002 (A.E. Kyles, BVMS, PhD, unpublished observations). Ninety-seven
percent of feline ureteral calculi contained calcium oxalate (A.E. Kyles,
BVMS, PhD, unpublished observations). There has been a marked increase

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in the incidence of calcium oxalate urolithiasis over the last 20 years. At
the University of Minnesota Urolith Center, the percentage of feline uro-
liths composed of calcium oxalate increased from 1% to 53% and the per-
centage composed of struvite decreased from 78% to 39% between 1981
and 1997

The frequency of intraluminal ureteral obstruction in cats in which

ureteral calculi are not diagnosed also seems to be increasing (A.E. Kyles,
BVMS, PhD, unpublished observation). Many cases respond to ﬂuid and
diuretic therapy; thus, a speciﬁc diagnosis is not reached. We have removed
soft tissue plugs, sometimes containing ﬂakes of mineralized material, and
dried solidiﬁed blood clots from obstructed feline ureters; however, bilateral
ureteral obstruction caused by blood clots has also been reported

Fibrosis has also been reported as a cause of ureteral obstruction in cats:
bilateral hydronephrosis caused by severe ureteral ﬁbrosis without an
obvious initiating cause has been reported in one cat

, and retroperito-

neal ﬁbrosis causing ureteral obstruction was diagnosed in four feline renal
transplant recipients 6 weeks to 5 months after transplantation

. A

number of other causes of ureteral obstruction in cats have been reported,
including congenital stenosis

, acquired stricture

, neoplasia (eg,

retroperitoneal leiomyosarcoma)

[37]

, and surgical trauma (eg, inadvertent

ligation of the ureters during ovariohysterectomy)

Any surgical procedure performed on the ureter can result in obstruction

caused by a stricture at the surgery site. The small size of the feline ureter
may predispose to this complication. Ureteral obstruction has been reported
after implantation of the ureter into the bladder (ureteroneocystostomy) in
cats using a number of surgical techniques

[8,39,40]

. An extravesical

modiﬁed Lich-Gregoir technique produces the least degree of ureteral
obstruction in cats with undilated ureters and is the preferred technique in
feline renal transplant patients

. The degree of postoperative ureteral

obstruction observed after ureteroneocystostomy in cats with preexisting
ureteral obstruction and ureteral dilation may be less than in cats with
normal ureters, such as renal transplant recipients.

Clinical ﬁndings

Clinical ﬁndings reported in dogs with ureteral obstruction include

abnormalities in urination (eg, urinary incontinence, stranguria, dysuria,
pollakiuria, polyuria, hematuria), persistent urinary tract infection, abdom-
inal pain, vomiting, anorexia, depression or lethargy, emaciation, fever or
hypothermia, palpation of abdominal mass, vaginal discharge, and prosta-
tomegaly

[16,18,19,22–25,41–43]

. Abdominal pain was common in dogs

with calculi, whereas urinary incontinence was seen in dogs with polyps.

Clinical signs reported in cats with ureteral obstruction are commonly

nonspeciﬁc, such as reduced appetite, lethargy, and weight loss

. Clinical

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signs may also be referable to uremia, such as vomiting, polyuria, and
polydipsia, or directly to ureteral obstruction, such as stranguria, pollakiu-
ria, hematuria, and abdominal pain

Imaging

Imaging of the ureter is challenging using conventional radiographic

techniques

[5,6,13]

. The small size of this structure and the superimposi-

tion of larger soft tissue structures make it almost impossible to see. Radio-
paque calculi, if large enough, are visible on plain abdominal radiographs
(

), and hydroureter, if severe, may also be seen. Intravenous (IV)

excretory urography may aid in visualization of the dilated ureter proximal
to the site of obstruction, but opaciﬁcation of the aﬀected kidney and ureter
is often poor.

Percutaneous antegrade pyelography provides good visualization of the

renal pelvis and ureter and has been used to determine if obstruction is
present (

)

[17,26]

. The animal is heavily sedated or anesthetized. IV

ﬂuids are administered during and after the procedure. The animal is placed
in dorsal recumbency, and the skin over the region of the kidney is surgically
prepared. Under ultrasound guidance, a 3.5-in, 22-gauge (dogs) or 2.5-in,
25-gauge (cats) spinal needle is placed through the greater curvature of the
kidney into the renal pelvis. Urine is withdrawn until the renal pelvis is
reduced in size by one half. This urine can be submitted for bacteriologic
culture and cytologic examination. A volume of aqueous iodinated contrast
material equal to half the urine volume removed is infused into the pelvis of
the kidney under ultrasound guidance. Alternatively, multiple small boluses
of contrast media can be administered and the region repeatedly imaged
with ﬂuoroscopy. The needle is withdrawn, and lateromedial and ventro-
dorsal radiographs are obtained immediately and 15 minutes later.

Ultrasonographic imaging has markedly increased the ability to detect

small increases in the size of the renal pelvis and ureter (

. If an

Fig. 2. Survey lateral and ventrodorsal radiographs (A, B) and intravenous pyelogram (IVP)
(C, D) from a 4-year-old male, castrated, domestic shorthair cat with a 4-day history of
depression, inappetence, stranguria, and pollakiuria. The survey radiographs show a moderately
sized radiopaque calculus in the region of the left renal pelvis. On the lateral view, there is ill-
deﬁned mineralization superimposed over the caudal poles of the kidneys. A small radiopaque
calculus is present in the bladder. The IVP radiographs were obtained 35 minutes after
administration of an iodinated contrast medium. There is marked distention and enlargement of
the right renal diverticula and pelvis and proximal ureter. The right ureter abruptly narrows,
and the distal end has an irregular tortuous appearance. The left renal pelvis is distorted and
mildly dilated. The cat was diagnosed with a left renal calculus, right ureteral obstruction, and
cystic calculus. A stricture of the right proximal ureter was found at surgery. (Courtesy of the
Digital Image Library of the Veterinary Medical Teaching Hospital, University of California at
Davis, Davis, CA.)

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obvious cause of ureteral dilation cannot be found, measuring the renal
resistive index (RRI) of the kidney may be helpful in separating obstructive
from nonobstructive disease

[13,44]

. The RRI of the normal canine kidney is

0.55 to 0.72 if no sedatives are used and 0.32 to 0.57 if sedatives are used.
The normal RRI of sedated cats is 0.52 to 0.63. The RRI increases after

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

obstruction, but thresholds of 0.70 to 0.73 were not always diagnostic for
obstruction in dogs. When dogs with unilateral complete obstruction were
given mannitol (0.5 g/kg administered intravenously), the RRI decreased in
the normal kidneys to 0.49 to 0.58 but remained at 0.75 to 0.86 in the
obstructed kidneys

[44]

. Comparison of the change in RRI of the normal

kidney and that of the obstructed kidney may help to conﬁrm obstruction.

Helical CT excretory urography has been used to image the ureter of

normal dogs, and CT is the modality of choice for imaging urolithiasis in
many human hospitals

. In cats, the sensitivity of CT and ultrasonography

for the diagnosis of ureteral dilation is similar, but CT is superior in the
determination of the number and position of ureteral calculi (

) (A.E.

Kyles, BVMS, PhD, unpublished observation).

Renal scintigraphy has been used clinically to evaluate the obstructed

kidney and ureter

. Technetium Tc 99m diethylenetriamine pentaacetic

Fig. 3. Survey lateral radiograph (A) and antegrade pyelogram (B) from a 7-year-old male,
castrated, domestic shorthair cat with a 2-week history of stranguria, pollakiuria, and dysuria.
Antegrade pyelography was performed on the left kidney by ultrasound-guided placement of
a spinal needle in the renal pelvis, extraction of urine, and administration of a lesser volume of
iodinated contrast medium. The left renal diverticula and pelvis are moderately dilated, and
there are multiple luminal defects within the contrast medium in the renal pelvis. The proximal
left ureter is moderately dilated and tortuous, and there is a luminal defect located at the distal
end of the contrast-ﬁlled ureter. Note the presence of a rectal temperature probe and
electrocardiographic lead. The radiographic diagnosis was left renal calculi, a proximal left
ureteral calculus, and left-sided ureteral obstruction. At surgery, an ureterotomy was performed
to remove a calculus from the left proximal ureter. (Courtesy of the Digital Image Library of the
Veterinary Medical Teaching Hospital, University of California at Davis, Davis, CA.)

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Fig. 4. Sagittal ultrasonographic view of the right kidney (A) and right retroperitoneal area (B)
from a 4-year-old female, spayed, domestic shorthair cat with a 3-day history of anorexia,
vomiting, polyuria, and polydipsia. There is a large amount of subcapsular ﬂuid surrounding an
enlarged right kidney with a moderately dilated renal pelvis. The proximal ureter is dilated for
approximately 1.5 cm (

þ—þ). A mineralized intraluminal object with shadowing is present

within the ureter (x—x). The ureter distal to the object is mildly dilated. The ultrasonographic
diagnosis is a proximal right ureteral calculus with hydronephrosis and hydroureter. The cat
was treated with hemodialysis, and the right ureteral calculus was removed via an ureterotomy.
(Courtesy of the Digital Image Library of the Veterinary Medical Teaching Hospital, University
of California at Davis, Davis, CA.)

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acid (DTPA) scintigraphy can be used to measure the GFR of individual
kidneys. The GFR in the obstructed kidney is reduced, and scintigraphy
cannot predict the GFR after relief of the obstruction. Measurement of the
GFR of the contralateral kidney may assist in the decision to perform
a nephrectomy on the obstructed kidney. It has also been used to monitor
obstruction after ureteroneocystostomy

[45]

. The technique is under

investigation for use in the diagnosis of ureteral obstruction in dogs, but
a clinically useful protocol has not yet been determined

. Ureteral

obstruction results in an increase in the renal transit time (deﬁned as the
time to peak

99m

Tc-DTPA activity) because of decreased renal blood ﬂow

Fig. 5. CT study of a 9-year-old male, castrated, domestic shorthair cat with a 1-week history of
lethargy and vomiting. Transverse 5-mm sections of the abdomen were obtained before (A) and
after (B) intravenous (IV) administration of iodinated contrast medium. The survey CT slice
shows radiopaque calculi in the left kidney and proximal ureter (arrow). The contrast CT slice
shows moderate dilation of the left renal pelvis. The diameter of the left ureter appears normal,
and the radiopaque calculus can still be observed (arrow). The cat was treated with IV ﬂuids and
diuretic drugs, and the serum creatinine concentration decreased from 7.9 mg/dL at
presentation to 2.2 mg/dL at recheck examination 8 weeks later. (Courtesy of the Digital
Image Library of the Veterinary Medical Teaching Hospital, University of California at Davis,
Davis, CA.)

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and GFR and an increase in the volume of the collecting system. Increased
whole-kidney transit time can be associated with dilation of the renal pelvis
without obstruction, however. Parenchymal transit time, calculated by
drawing the region of interest to exclude the collecting system, is a more
reliable indicator of renal obstruction.

Medical and minimally invasive treatment

Animals with ureteral obstruction often present in renal failure and must

be stabilized before undergoing invasive treatment to remove the obstruc-
tion. Fluid diuresis alone or in combination with diuretic drug therapy may
result in relief of intraluminal causes of obstruction

. In cats, there is

anecdotal evidence that glucagon (0.1 mg per cat administered intravenously
twice daily) may cause relaxation of the ureteral smooth muscle and
promote passage of ureteral calculi. Severe azotemia should be treated
before deﬁnitive surgery with hemodialysis

if available or with drainage

of urine via a nephrostomy tube placed in the pelvis of the obstructed kidney

[38,48]

. Nephrostomy tubes can be placed percutaneously under ultrasound

guidance; in cats, we have percutaneously placed 4-French Swan-Ganz or 5-
French Dawson-Mueller catheters. Alternatively, a nephrostomy catheter
can be placed via an emergency laparotomy; either a 5-French red rubber or
5-French Foley catheter can be used in cats. Nephrostomy tube placement
requires that the renal pelvis is dilated and so may not be possible in animals
with acute obstruction. Preoperative nephrostomy tube placement has the
additional advantage of allowing an evaluation of the remaining function in
the obstructed kidney before deﬁnitive surgery.

Extracorporeal shock-wave lithotripsy using dry and water bath techni-

ques has been used to fragment ureteroliths in the dog

[41,49,50]

. Several

treatments may be required. Lithotripsy is not currently recommended for
cats, because the feline kidney is more sensitive to shock wave–induced
injury

. In human beings, ureteroscopy is also used to remove calculi

, but this technique has not been adapted for the dog and cat. Overall,

the move to noninvasive treatment of urolithiasis in human beings has
improved renal preservation by a factor of 10

[52]

Surgical treatment

If obstruction is unilateral and there is adequate function in the opposite

kidney, unilateral removal of the kidney and ureter is an option for
treatment of ureteral obstruction

[2,42,48]

. The obstruction may be bi-

lateral, the animal may be at risk for recurrence of obstruction in the
opposite ureter, or renal function in the opposite kidney may be impaired,
however. In these instances, preservation of renal function on the aﬀected
side is critical, and ureteral surgery is necessary.

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Healing in the ureter occurs by ﬁbrous tissue replacement. When 5-cm

ureteral defects were created in dogs and healing occurred over a stent, the
healed ureter was lined with uroepithelium but was narrow and the wall
consisted mainly of ﬁbrous tissue

[53]

. Given the small size of the ureter,

particularly in the cat, and the need for precise surgical technique, surgeons
attempting ureteral surgery should be well versed in microsurgical techni-
ques. The use of surgical loupes or, particularly in cats, an operating
microscope greatly facilitates surgery.

Surgical removal of ureteral calculi or other causes of intraluminal

ureteral obstruction is indicated when a ureter is partially or completely
obstructed as indicated by hydronephrosis and hydroureter proximal to the
calculus and immobility of the ureteral calculus as determined by repeated
radiographic or ultrasonographic examinations

[29,54]

. Ureteral calculi can

be removed by ureterotomy or, when located in the distal ureter, by
ureteroneocystostomy. In cats, it has been recommended that ureteral
calculi in the proximal third of the ureter be removed by ureterotomy with
urine drainage via a nephrostomy catheter, whereas ureteral calculi in the
distal two thirds of the ureter can be managed by partial ureterectomy and
ureteroneocystostomy

. In animals with ureteral obstruction caused by

mural lesions or extraluminal obstruction, ureteroneocystostomy or ureteral
resection and anastomosis (ureteroureterostomy) may be needed to re-
construct the ureter and preserve ipsilateral renal function.

Ureterotomy

The primary indication for ureterotomy is the removal of ureteral calculi

or other causes of intraluminal ureteral obstruction (

Figs. 6, 7

)

[2,4,29]

. The

ureter is approached via a midline laparotomy. Ureteral calculi may be
directly observed or palpated. Ureteral dilation tends to begin proximally
and extend distally so that the dilated ureter may not extend all the way to
the level of the calculus. If a calculus cannot be identiﬁed, a cystotomy
should be performed and the ureter should be catheterized from the bladder.
The ureter should be manipulated carefully, and the ureteral blood supply
should be preserved. The aﬀected portion of the ureter is mobilized from the
retroperitoneal space. In cats, the ureter is surrounded by fat, which needs to
be dissected away from the ureter. A transverse or longitudinal incision is
made with a scalpel blade in the dilated ureter proximal to the ureteral
calculi. We prefer a longitudinal incision, which can then be extended using
scissors. Normally, the calculus is readily removed, but it occasionally
becomes embedded in the ureteral wall. Care should be taken to prevent
a ureteral calculus being pushed back into the renal pelvis during
manipulation of the ureter. After the calculus is removed, the ureter is
ﬂushed with warm saline using a lacrimal duct needle and a catheter or, in
cats, a length of suture material passed proximally into the renal pelvis and
distally into the bladder to ensure that no additional calculi remain. The

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ureterotomy incision is closed with 5-0 to 8-0 monoﬁlament suture material.
We prefer a simple continuous suture pattern using a full-thickness bite to
ensure a watertight seal.

Ureteroneocystostomy

Ureteroneocystostomy involves implantation of the distal ureter into the

bladder. The procedure is indicated to re-establish urine ﬂow after resection
of distal ureteral lesions and is also used during renal transplantation and in
cases of extramural ureteral ectopia and ureteral avulsion. Surgical
techniques for ureteroneocystostomy can be divided into intravesical
techniques, such as the mucosal apposition technique

, which are

performed from within the bladder lumen and require a cystotomy for

Fig. 6. Ureterotomy for the removal of obstructive ureteral calculi. (A) Ureteral calculus lodged
within the lumen of the ureter. Ureteral dilation occurs proximal to the site of obstruction.
Several methods of ureterotomy are considered acceptable. (B) A longitudinal ureterotomy
incision is made in the dilated segment proximal to the calculus. The calculus is removed, and
the incision is closed longitudinally. (C) For preservation of luminal diameter, the longitudinal
ureterotomy incision can be closed transversely. (D) A transverse ureterotomy incision is closed
in a transverse manner after calculus removal. (From McLouglin MA, Bjorling DE. Ureters. In:
Slatter D, editor. Textbook of small animal surgery. 3rd edition. Philadelphia: WB Saunders;
2003. p. 1619–28; with permission.)

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access, and extravesical techniques, such as the modiﬁed Lich-Gregoir
technique

. The mucosal apposition technique works well in dogs, but

the modiﬁed Lich-Gregoir technique is preferred in cats, particularly when
there is minimal preexisting ureteral dilation, because it is associated with
a reduced degree of postoperative swelling and ureteral obstruction

The mucosal apposition technique requires a ventral cystotomy and

eversion of the bladder (

). A stab incision is made in the bladder

mucosa, and a short oblique tunnel is created by pushing microhemostats
through the bladder wall from inside to outside. The distal end of the
transected ureter is grasped with the hemostats, and the end of the ureter is
pulled into the bladder lumen. The distal ureteral vessels should be identiﬁed
and ligated if possible; electrocautery should not be used on the ureter.
Periureteral fat is dissected from the ureter for a distance of 0.5 to 1 cm,
preserving the blood vessels; this is particularly important in cats, in which
there is a considerable amount of periureteral fat. The ureter is cut
longitudinally to create a 0.5-cm section of spatulation. The mucosa of
the spatulated ureter is sutured to the bladder mucosa using a simple
interrupted suture pattern and 6-0 to 8-0 monoﬁlament suture material. The
ﬁrst suture should be placed at the apex of the spatulation. A catheter or, in
cats, length of suture material should be placed in the ureter to ensure
patency. The second suture is placed at the distal end of the spatulation, and
four to six additional sutures are then placed. There should be accurate
mucosa-to-mucosa apposition. In cats, the bladder mucosa rapidly becomes
edematous, which makes suturing more challenging and may contribute to
a degree of postoperative ureteral obstruction.

Fig. 7. Closure of an ureterotomy incision in a cat. An operating microscope greatly facilitates
this surgery. The incision is closed with 8-0 monoﬁlament suture material. A suture is placed,
and a knot is tied at one end of the incision, leaving a long end. A simple continuous suture is
placed, starting at the opposite end of the incision. Full-thickness bites should be taken. Tying
the continuous suture line to the single simple interrupted suture completes the suture pattern.
(Courtesy of Helen Mehl, MD, Koloa, HI; Margo Mehl, DVM, Davis, CA; and John Doval,
BPA, Sacramento, CA.)

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

Fig. 8. Removal of calculus from proximal ureter by intravesical ureteroneocystostomy. (A)
Proximal ureteral obstruction caused by calculus. (B) Nephrocystopexy to decrease distance
between the bladder and kidney and tension on anastomosis. (C) Performance of ureter-
oneocystostomy. (D) Completed surgery. (From Rawlings CA, Bjorling DE, Christie BA.
Kidneys. In: Slatter D, editor. Textbook of small animal surgery. 3rd edition. Philadelphia: WB
Saunders; 2003. p. 1606–19; with permission.)

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

The modiﬁed Lich-Gregoir technique does not require a cystotomy for

access (

Fig. 9

). A 1-cm seromuscular incision is made at the apex of the

bladder. The mucosa will bulge through this incision. A 0.5-cm incision is
made through the mucosa at the caudal aspect of the seromuscular incision.
The ureter is spatulated as described previously. The mucosa of the spatulated
ureter is sutured to the bladder mucosa in a similar manner to that of the
mucosal apposition technique. A minimum number of simple interrupted
sutures, usually six, are required, because an excessive number increase the
risk of postoperative ureteral obstruction. The seromuscular layer is closed
with simple interrupted sutures using 5-0 monoﬁlament suture material,
taking care not to compress the ureter caudally. The seromuscular layer rather
than the mucosal layer provides the watertight seal.

Ureteroureterostomy

Ureteroureterostomy is indicated to repair the ureter after ureteral

resection or transection (

Fig. 10

). The procedure is technically more diﬃcult

than ureteroneocystostomy and is associated with a higher incidence of
postoperative ureteral obstruction. Hence, ureteroureterostomy is normally
performed only when the proximal end of the ureter cannot be implanted
into the bladder directly. It is also possible to attach the ureter to the
contralateral ureter (transureteroureterostomy)

[56]

The chances of ureteral stricture after ureteroureterostomy are reduced

by increasing the circumference of the anastomosis. This can be accom-
plished by spatulation or by transecting the ureter obliquely. The spat-
ulation technique involves longitudinal incisions in both ureteral ends,
which should be positioned on opposite sides of the ureter. The anastomosis
is performed using a simple interrupted suture pattern and 6-0 to 8-0
monoﬁlament suture material. The ﬁrst two sutures placed connect the
apex of the spatulation to the corresponding portion of the opposite end of
the ureter, taking care not to twist the ureteral ends. Simple interrupted
sutures are then placed to complete the anastomosis.

Techniques for tension reduction

Tension on the ureteroneocystostomy or ureteroureterostomy anastomo-

sis should be avoided. Tension can be reduced by performing renal
descensus and psoas cystopexy

[29,57]

. Renal descensus involves dissecting

the kidney from its peritoneal attachments so that it remains attached by the
renal vessels, repositioning the kidney more caudally, and performing
a nephropexy to attach the kidney to the body wall and prevent renal
torsion. Psoas cystopexy involves pulling the apex of the bladder cranially
and laterally toward the kidney and suturing the bladder to the psoas muscle
dorsally. An alternative technique for reducing the tension on the
anastomosis is nephrocystopexy. This involves dissecting the kidney from

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

Fig. 9. Modiﬁed Lich-Gregoir technique for extravesical ureteroneocystostomy used in cats.
(A, B) A seromuscular incision is made at the apex of the bladder. An incision approximately
half the length of the seromuscular incision is made through the mucosa at the caudal aspect of
the seromuscular incision. The excess fat is dissected from the distal ureter. A longitudinal
incision is made in the ureter. The mucosa of the spatulated ureter is sutured to the bladder
mucosa. The ﬁrst two sutures of 8-0 monoﬁlament suture material are placed cranially at the
apex of the spatulation and caudally at the distal end of the ureter. (C) Four additional simple
interrupted sutures are placed. (D) The seromuscular layer is closed with simple interrupted
sutures using 5-0 monoﬁlament suture material, taking care not to compress the ureter cranially.
Note that the seromuscular layer rather than the mucosal layer provides the watertight seal.
(From Bernsteen L, Gregory CR, Kyles AE, et al. Renal transplantation in cats. Clin Tech
Small Anim Pract 2000;15:40–5; with permission.)

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

its retroperitoneal attachments, repositioning it caudally, pulling the bladder
forward, and suturing the kidney directly to the bladder.

Urine diversion

Urine can be diverted after ureterotomy or ureteroureterostomy by

placement of a ureteral stent or nephrostomy tube. Ureteral stents can be
placed for 3 to 10 days for urine diversion and then recovered by
cystoscopy. Alternatively, the stent can be exteriorized via a cystostomy
and hooked up to a closed drainage system, which allows quantiﬁcation of
urine output and easy removal of the ureteral stent

[58,59]

. Ureteral stents

are controversial, however, and the presence of a stent across the surgical
site may increase ﬁbrosis and the risk of ureteral stricture. Nephrostomy
tube placement is an alternative method of urine diversion

[29,38]

. After

a proximal ureterotomy, a nephrostomy tube can be placed using a 20-
gauge over-the-needle IV catheter inserted through the ureterotomy in-
cision, advanced through the renal pelvis, and exited through the renal
cortex

. A length of suture material is tied to a 5-French red rubber or

Foley catheter, and the other end of the suture is passed through the IV
catheter. The catheter and suture are slowly withdrawn, pulling the tip of

Fig. 10. Ureteroureterostomy (ureteral anastomosis). (A) The proximal and distal ureteral ends
are debrided and incised longitudinally to widen each ureteral oriﬁce and to provide a longer
circumference of the ureteral oriﬁce for suturing. (B) The anastomosis is performed with 5-0 to
8-0 monoﬁlament suture material. Two sutures are placed 180( apart at the apex of each
longitudinal incision. (C) A continuous or interrupted suture pattern is used to complete the
anastomosis. (From McLouglin MA, Bjorling DE. Ureters. In: Slatter D, editor. Textbook
of small animal surgery. 3rd edition. Philadelphia: WB Saunders; 2003. p. 1619–28; with
permission.)

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

the nephrostomy tube into the ureterotomy incision. The suture is then cut,
and the tip of the catheter is positioned in the renal pelvis. A nephrostomy
tube can be placed in patients without a proximal ureterotomy incision by
placing a 16-gauge over-the-needle IV catheter through the body wall from
lateral to medial adjacent to the kidney and then through the kidney into
the renal pelvis

. A 3.5-French red rubber catheter with the tip cut oﬀ is

passed down the IV catheter, and the tip is positioned in the renal pelvis.
The IV catheter is withdrawn and seated onto the hub of the red rubber
catheter. With either technique, a nephropexy is performed by placing
sutures between the kidney and body wall around the nephrostomy tube.
The nephrostomy tube is secured to the skin and connected to a closed
drainage system.

In cats, the undilated ureter is too small to allow placement of a ureteral

stent, and nephrostomy tubes are associated with a high complication rate
with problems of poor drainage, tube dislodgement, and urine leakage.
Tube dislodgement is the most common problem and is frequently
associated with a nephrostomy tube that is attached to the skin, where the
mobility of the skin allows the tube tip to migrate. The chances of tube
dislodgement can be minimized by securing the tube to the outside of the cat
with a deep suture that incorporates the skin and body wall. It is our
experience that ureterotomy incisions in cats can heal satisfactorily without
urine drainage via a nephrostomy tube.

Prognosis

The prognosis for recovery of renal function after relief of ureteral

obstruction depends on a number of factors, including the duration and
degree of obstruction. In experimental dogs, 4 days of complete obstruction
resulted in near-complete return of renal function with correction, 14 days
of obstruction resulted in a 46% recovery in GFR and tubular function by
4 months after correction, and 40 days of obstruction resulted in little
recovery of renal function after correction

[60–62]

. Resolution of hydro-

ureter with time has been reported after surgery in dogs with naturally
occurring ureteral obstruction

[58,59,63]

. The success rate for lithotripsy in

32 dogs with renal or ureteral calculi was 90%, with 30% of dogs requiring
two or more treatments

In cats with ureteral calculi, the rates of major postoperative complica-

tions and perioperative mortality with surgical treatment are 30% and 19%,
respectively (A.E. Kyles, BVMS, PhD, unpublished observations). The most
common postoperative complications are urine leakage and ureteral
obstruction. Long-term follow-up after successful surgical removal of
ureteral calculi shows that surgery improved or preserved renal function

www.vetsurgerycentral.com/

. The rate of recurrence of ureteral calculi after removal is unknown.

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E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

Summary

The most common cause of ureteral obstruction in dogs and cats is

ureteral calculi. Common clinical signs associated with ureteral obstruction
include abnormalities in urination, persistent urinary tract infection,
abdominal pain, vomiting, anorexia, weight loss, and depression or lethargy.
Medical management of ureteral obstruction includes ﬂuid diuresis, muscle
relaxants, and treatment of azotemia using nephrostomy tubes or hemodi-
alysis. Surgical techniques used to restore patency to the ureter include
ureterotomy, partial ureterectomy and ureteroneocystostomy, and ureteral
resection and anastomosis. Lithotripsy has been used in dogs to remove
ureteral calculi. Renal function can be preserved if complete ureteral
obstruction is relieved within several days of onset.

References

[1] Christie BA. Anatomy of the urinary tract. In: Slatter D, editor. Textbook of small animal

surgery. 3rd edition. Philadelphia: WB Saunders; 2003. p. 1558–75.

[2] Lanz OI, Waldron DR. Renal and ureteral surgery in dogs. Clin Tech Small Anim Pract

2000;15:1–10.

[3] Weiss RM. Physiology and pharmacology of the renal pelvis and ureter. In: Retik AB,

Vaughan ED Jr, Wein AJ, editors. Campbell’s urology. 8th edition. Philadelphia: WB
Saunders; 2002. p. 377–409.

[4] Degner DA, Mathews K. Microureteral surgery. Available at:

ureteral_anatomy.htm

. Accessed October, 2003.

[5] McLouglin MA, Bjorling DE. Ureters. In: Slatter D, editor. Textbook of small animal

surgery. 3rd edition. Philadelphia: WB Saunders; 2003. p. 1619–28.

[6] Rozear L, Tidwell AS. Evaluation of the ureter and ureterovesicular junction using helical

computed tomographic excretory urography in healthy dogs. Vet Radiol Ultrasound 2003;
44:155–64.

[7] Kabalin JN. Surgical anatomy of the retroperitoneum, kidneys, and ureters. In: Retik AB,

Vaughan ED Jr, Wein AJ, editors. Campbell’s urology. 8th edition. Philadelphia: WB
Saunders; 2002. p. 3–40.

[8] Kochin EJ, Gregory CR, Wisner E, et al. Evaluation of a method of ureteroneocystostomy

in cats. J Am Vet Med Assoc 1993;202:257–60.

[9] Wen JG, Frokiaer J, Jorgensen TM, et al. Obstructive nephropathy: an update of the

experimental research. Urol Res 1999;27:29–39.

[10] Gulmi FA, Felsen FA, Vaughan ED Jr. Pathophysiology of urinary tract obstruction. In:

Retik AB, Vaughan ED Jr, Wein AJ, editors. Campbell’s urology. 8th edition.
Philadelphia: WB Saunders; 2002. p. 411–56.

[11] Chuang YH, Chuang WL, Liu KM, et al. Tissue damage and regeneration of ureteric

smooth muscle in rats with obstructive uropathy. Br J Urol 1998;82:261–6.

[12] Pullium JK, Dillehay DL, Webb S, et al. Congenital bilateral ureteral stenosis and

hydronephrosis in a neonatal puppy. Contemp Top Lab Anim Sci 2000;39:34–6.

[13] Lamb CR. Ultrasonography of the ureters. Vet Clin North Am Small Anim Pract 1998;28:

823–48.

[14] Franti CE, Ling GV, Ruby AL, et al. Urolithiasis in dogs. V: regional comparisons of breed,

age, sex, anatomic location, and mineral type of calculus. Am J Vet Res 1999;60:29–42.

[15] Armbrust L, Kraft SL, Cowan LA, et al. Radiographic diagnosis: canine ureteral calculus.

Vet Radiol Ultrasound 1997;38:360–2.

1008

E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

[16] Dupre GP, Dee LG, Dee JF. Ureterotomies for treatment of ureterolithiasis in two dogs.

J Am Anim Hosp Assoc 1990;26:500–4.

[17] Rivers BJ, Walter PA, Polzin DJ. Ultrasonographic-guided, percutaneous antegrade

pyelography: technique and clinical application in the dog and cat. J Am Anim Hosp Assoc
1997;33:61–8.

[18] Lautzenhiser SJ, Bjorling DE. Urinary incontinence in a dog with ectopic ureterocele. J Am

Anim Hosp Assoc 2002;38:29–32.

[19] Stiﬄer KS, Stevenson MA, Mahaﬀey MB, et al. Intravesical ureterocele with concurrent

renal dysfunction in a dog: a case report and proposed classiﬁcation system. J Am Anim
Hosp Assoc 2002;38:33–9.

[20] van Kempen M, Eble MJ, Krempien R, et al. Inﬂuence of irradiated volume on ureteral

injury after intraoperative radiation therapy: experimental study in dogs. Radiology 2003;
228:139–43.

[21] Burton CA, Day MJ, Hotston Moore A, et al. Ureteric ﬁbroepithelial polyps in two dogs.

J Small Anim Pract 1994;35:593–6.

[22] Reichle JK, Peterson RA II, Mahaﬀey MB, et al. Ureteral ﬁbroepithelial polyps in four

dogs. Vet Radiol Ultrasound 2003;44:433–7.

[23] Crawford MA, Turk MAM. Ureteral obstruction associated with proliferative ureteritis in

a dog. J Am Vet Med Assoc 1984;184:586–8.

[24] de Gopegui R, Espada Y, Majo N. Bilateral hydroureter and hydronephrosis in a nine-

year-old female German shepherd dog. J Small Anim Pract 1999;40:224–6.

[25] McEvoy FJ. Iatrogenic renal obstruction in a dog. Vet Rec 1994;135:457–8.
[26] Adin CA, Herrgesell EJ, Nyland TG, et al. Antegrade urography in cats with suspected

ureteral obstruction: 11 cases (1995–2001). J Am Vet Med Assoc 2003;222:1576–81.

[27] Crowell WA, Cornelius LM, Barsanti JA, et al. Unusual oliguric urinary failure in a cat.

J Am Anim Hosp Assoc 1982;18:812–4.

[28] Gotthelf LN. Three concurrent urinary diseases in a cat: urinary obstruction, hypoplasia of

the kidney and nephritis caused by feline infectious peritonitis. Vet Med Small Anim Clin
1981;76:217–9.

[29] Kyles AE, Stone AE, Gookin J, et al. Diagnosis and surgical management of obstructive

ureteral calculi in cats: 11 cases (1993–1996). J Am Vet Med Assoc 1998;213:1150–6.

[30] Moon ML, Dallman MA. Calcium oxalate urolith in a cat. Vet Radiol 1991;32:261–3.
[31] Wolf AM, Leighton RL, Watrous BJ. Uric acid ureteral calculi and pararenal cyst in a cat.

J Am Anim Hosp Assoc 1979;15:767–73.

[32] Lekcharoensuk C, Lulich JP, Osborne CA, et al. Association between patient-related

factors and risk of calcium oxalate and magnesium ammonium phosphate urolithiasis in
cats. J Am Vet Med Assoc 2002;217:520–5.

[33] Lieb MS, Allen TA, Konde LJ, et al. Bilateral hydronephrosis attributable to bilateral

ureteral ﬁbrosis in a cat. J Am Vet Med Assoc 1988;192:795–7.

[34] Aronson LR. Retroperitoneal ﬁbrosis in four cats following renal transplantation. J Am

Vet Med Assoc 2002;221:984–9.

[35] North DC. Hydronephrosis and hydro-ureter in a kitten—a case report. J Small Anim

Pract 1978;19:237–40.

[36] Prasad MC, Mohan K. Hydronephrosis in the cat. Indian Vet J 1968;45:107–8.
[37] Speakman CF, Pechman RD, D’Andrea GH. Aortic thrombosis and unilateral hydro-

nephrosis associated with leiomyosarcoma in a cat. J Am Vet Med Assoc 1983;182:62–3.

[38] Nwadike BS, Wilson LP, Stone EA. Use of bilateral temporary nephrostomy catheters for

emergency treatment of bilateral ureter transaction in a cat. J Am Vet Med Assoc 2000;
217:1862–5.

[39] Gregory CR, Lirtzman RA, Kochin EJ, et al. A mucosal apposition technique for

ureteroneocystostomy after renal transplantation in cats. Vet Surg 1996;25:13–7.

[40] Mehl ML, Kyles AE, Pollard R, et al. Comparison of three ureteroneocystostomy

techniques in cats [abstract]. Vet Surg 2002;31:490–1.

1009

E.M. Hardie, A.E. Kyles / Vet Clin Small Anim 34 (2004) 989–1010

[41] Bailey G, Burk RL. Dry extracorporeal shock wave lithotripsy for treatment of

ureterolithiasis in a dog. J Am Vet Med Assoc 1995;207:592–5.

[42] Gookin JL, Stone EA, Spaulding KA, et al. Unilateral nephrectomy in dogs with renal

disease: 30 cases (1985–1994). J Am Vet Med Assoc 1996;208:2020–6.

[43] Hattel AL, Diters RW, Snavely DA. Ureteral ﬁbropapilloma in a dog [abstract]. J Am Vet

Med Assoc 1986;188:873.

[44] Choi H, Chung W, Lee K, et al. Eﬀect of intravenous mannitol upon the resistive index in

complete unilateral renal obstruction in the dog. J Vet Intern Med 2003;17:158–62.

[45] Barthez PY, Smeak DD, Wisner ER, et al. Ureteral obstruction after ureteroneocystos-

tomy in dogs assessed by technetium TC 99m diethylenetriamine pentaacetic acid (DTPA)
scintigraphy. Vet Surg 2000;29:499–506.

[46] Barthez PY, Smeak DD, Wisner ER, et al. Eﬀect of partial ureteral obstruction on results

of renal scintigraphy in dogs. Am J Vet Res 1999;60:1383–9.

[47] Langston CE, Cowgill LD, Spano JA. Applications and outcome of hemodialysis in cats:

a review of 29 cases. J Vet Intern Med 1997;11:348–55.

[48] Rawlings CA, Bjorling DE, Christie BA. Kidneys. In: Slatter D, editor. Textbook of small

animal surgery. 3rd edition. Philadelphia: WB Saunders; 2003. p. 1606–19.

[49] Adams LG, Senior DF. Electrohydraulic and extracorporeal shock-wave lithotripsy. Vet

Clin North Am 1999;29:293–302.

[50] Block G, Adams GA, Widmer WR, et al. Use of extracorporeal shock wave lithotripsy for

treatment of nephrolithiasis and ureterolithiasis in ﬁve dogs. J Am Vet Med Assoc 1996;
208:531–6.

[51] Gettman LY, Roehrborn CG, Cadeddu JA, et al. Management of ureteral calculi: a cost

comparison and decision making analysis. J Urol 2002;167:1621–9.

[52] Holman CD, Wisniewski ZS, Semmens JB, et al. Changing treatments for primary

urolithiasis: impact on services and renal preservation on 16,679 patients in Western
Australia. Br J Urol Int 2002;90:7–15.

[53] Kuzaka B, Szymanska K, Borkowski A, et al. Restoration of the continuity of dog ureter

after resection of its 5 cm middle segment. Br J Urol 1996;77:342–6.

[54] Caywood DD, Osborne CA. Surgical removal of canine uroliths. Vet Clin North Am 1986;

16:389–407.

[55] Bernsteen L, Gregory CR, Kyles AE, et al. Renal transplantation in cats. Clin Tech Small

Anim Pract 2000;15:40–5.

[56] Crane SW, Waldron DR. Ureteral function and healing following microsurgical

transureteroureterostomy in the dog. Vet Surg 1980;9:108–15.

[57] Stone EA, Barsanti JA. Surgical therapy for urinary tract trauma. In: Urologic surgery of

the dog and cat Philadelphia: Lea & Febiger; 1992. p. 189–98.

[58] Chambers JN, Selcer BA, Barsanti JA. Recovery from severe hydroureter and

hydronephrosis after ureteral anastomosis in a dog. J Am Vet Med Assoc 1987;191:
1589–92.

[59] Mehl ML, Kyles AE. Ureteroureterostomy after proximal ureteral injury during

ovariohysterectomy in a dog. Vet Rec 2003;153:469–70.

[60] Fink RL, Caridis DT, Chimile R, et al. Renal impairment and its reversibility following

variable periods of complete ureteral obstruction. Aust NZ J Surg 1980;50:77–83.

[61] Vaughan ED Jr, Gillenwater JY. Recovery following complete chronic unilateral

obstruction: functional, radiographic, and pathologic alterations. J Urol 1971;106:27–35.

[62] Vaughan ED Jr, Sweet RE, Gillenwater JY. Unilateral ureteral occlusion: pattern of

nephron repair and compensatory response. J Urol 1973;109:979–82.

[63] Ross LA, Lamb CR. Reduction of hydronephrosis and hydroureter associated with ectopic

ureters in two dogs after ureterovesical anastomosis. J Am Vet Med Assoc 1990;196:
1497–9.

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Lithotripsy: an update on urologic

applications in small animals

India F. Lane, DVM, MS

Department of Small Animal Clinical Sciences, University of Tennessee,

College of Veterinary Medicine, C247 Veterinary Teaching Hospital, 2407 River Drive,

Knoxville, TN 37996–4544, USA

Management of upper tract urolithiasis in human patients has undergone

dramatic transformation since 1980, when the ﬁrst clinical lithotriptor was
introduced by Christian Chaussy and collaborators

at Dornier Aero-

space Engineering. In the ensuing quarter century, extracorporeal, percuta-
neous, and ureteroscopic methods have largely replaced surgical intervention
for renal or ureteral stones. In veterinary medicine, similar technologic
advances have provided noninvasive means for reduction or elimination of
upper and lower tract stones. The fundamental principles of shock wave
methods, including extracorporeal shock wave lithotripsy (ESWL) and
intracorporeal electrohydraulic shock wave lithotripsy (EHL), have been
reviewed

. This article provides an update on extracorporeal methods and

introduces intracorporeal laser lithotripsy methods, which may be increas-
ingly available for small animal applications in the future.

Extracorporeal shock wave lithotripsy technology

Successful lithotripsy requires a source to generate shock waves (SWs),

a method for focusing the SWs to a solitary point, and a method for
transmitting (‘‘coupling’’) the SWs to the patient (

). SWs are high-

amplitude sound waves generated by electrohydraulic, electromagnetic, or
piezoelectrical energy sources. Like ultrasound waves, SWs travel through
media of ﬂuid or soft tissue density until reaching the ‘‘hard’’ acoustic
surface of the urolith. Energy reﬂection and creation of tensile stresses along
the surfaces of the stone as well as generation of cavitation bubbles within
the stone lead to multiple coalescing cracks (spallation) and fragmentation
with repeated SWs

. Original EWSL methods relied on transmission of

E-mail address:

ilane@utk.edu

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.013

Vet Clin Small Anim

34 (2004) 1011–1025

Table 1
Features of extracorporeal lithotriptors employed in veterinary patients

Model

Manu-

Source

Focusing

Coupling

Imaging

Focal Zone

Peak Pressure

HM-3

Dornier

EHL

Ellipsoid

Water bath

Radiograph

90 mm

1300 bar

Modulith

Storz

Electromagnetic

Parabolic

Water cushion

Rad/US

30 mm

1000 bar

MFL-5000

Dornier

EHL

Ellipsoid

Water cushion

Rad/US

40 mm

1000 bar

Piezolith 2500

Wolf

Piezoelectric

Concave dish

Water cushion

Rad/US

1.5

11 mm

1200 bar

Lithostar II

Siemens

Electromagnetic

Acoustic lens

Water cushion

Rad/US

80 mm

500–800 bar

Abbreviations

: EHL, electrohydraulic; Rad/US, both radiographic (ﬂuoroscopic) and ultrasonographic imaging modalities available.

1012

I.F.

Lane

Vet

Clin

Small

Anim

34
(2004)

1011–10

SWs through a water bath medium, requiring the patient to be partially
submerged during treatment. SWs were generated by the pulsatile sparks
created by an electrohydraulic electrode

. Newer lithotriptors use other

SW generators and ‘‘dry’’ methods, in which SWs are coupled to the patient
through a ﬂuid-ﬁlled cushion while the patient lies on an adjacent plastic
cradle (

Figs. 1 and 2

)

[5,6]

. Dual-imaging capabilities with ﬂuoroscopic and

ultrasonographic tracking of uroliths and variable power settings are
additional features of newer lithotriptors

. Although these lithotriptors

are easier to use and to maintain, the eﬃcacy of dry lithotriptors is lower
than that of the ‘‘gold standard’’ water bath model because of a smaller
focal zone and lower peak pressure. This narrow focal zone makes accurate
targeting of uroliths imperative; however, it also limits SW damage to
surrounding tissues

. The latest (so-called ‘‘third’’) generation of

lithotriptors is designed to increase portability and ﬂexibility in treatment
method. Machines with movable SW application sources may be useful for
reaching uroliths in diﬃcult locations and may allow nonurologic applica-
tions (eg, orthopedic) to be delivered by the same lithotriptor. Machines are
also designed to be smaller and less costly than prior prototypes

. Most of

these lithotriptors sacriﬁce eﬃciency and depth of penetration, however,
which may limit their eﬀectiveness for nephroliths in larger patients.

Fig. 1. Dry lithotripter unit with integrated C-arm ﬂuoroscopic capability, movable treatment
table, x-ray generator, and inline ultrasound capability.

1013

I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

Method for ‘‘dry’’ extracorporeal shock wave lithotripsy

At the University of Tennessee College of Veterinary Medicine, we are

using a Storz Modulith SL-20 ‘‘dry’’ lithotriptor (see

; Karl Storz,

Atlanta, GA). This lithotriptor employs an electromagnetic energy source,
a cylindric parabolic focusing reﬂector, and a water cushion to transmit (or
‘‘couple’’) the SWs to the patient. The patient rests on a ﬂexible acoustic
cradle, and the table can be moved from a focusing position under an
attached C-arm to a treatment position over the SW generator. An inline
ultrasound transducer also aids in isolating nephroliths and can be used for
real-time monitoring of stone fragmentation. We have found it diﬃcult to
use the inline ultrasound imaging in dogs and cats, however.

Candidates are evaluated with a routine database, urine culture, co-

agulation proﬁle, blood pressure measurement, abdominal radiographs,
abdominal ultrasound, renal scintigraphy, and, possibly, CT. Treatments
are completed under general anesthesia for optimal focusing control and
analgesia.

Fluoroscopy and ultrasound are used to localize the stone to be treated;

the patient is then placed over a soft cushion ﬁlled with water. SWs are
transmitted through the cushion to the treatment site at increasing power
levels (

). With this technique, there is no need for the patient to be

submerged in water; however, overlying fur is shaved, and water or gel is
used under the patient to facilitate SW transmission. Up to 2000 or 3000
SWs may be administered at one site. Progress is monitored periodically by
repeating ﬂuoroscopic imaging or using spot ﬁlms. Treatment time is usually
approximately 1 hour or longer if multiple sites are treated. Total anesthesia
time ranges from 1.5 to 2 hours.

All patients remain in the hospital 2 nights after treatment for ﬂuid

diuresis and initial follow-up radiographs, ultrasound, and repeat complete

Fig. 2. Coupling cushion for ‘‘dry’’ lithotripsy. The cushion is raised to contact the patient lying
on the overlying plastic cradle, and gel is used to facilitate transmission of shock waves from the
cushion surface to the patient.

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

blood cell count, serum chemistry panel, urinalysis, and, possibly, serum
amylase and lipase measurements. Patients are discharged with instructions
for monitoring urination and urine appearance, lethargy, apparent abdom-
inal pain, or vomiting. Small amounts of blood may be observed in the
urine, especially if they were seen before treatment; however, the blood
usually clears before discharge from the hospital. Follow-up chemistry
results and radiographs are recommended approximately 3 to 4 weeks after

Fig. 3. An example of a dog undergoing extracorporeal shock wave lithotripsy treatment. (A)
The urolith to be treated is positioned in the focal zone using ﬂuoroscopic guidance. (B) The
patient is returned to the treatment position for shock wave application.

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

treatment and then monthly for 2 to 3 months depending on the degree of
remaining urolith material. If we are concerned about ureteral obstruction
or urinary tract infection (UTI), follow-up ultrasound examination and
urine culture results are recommended as well. Long-term management is
recommended based on the known or ‘‘guesstimated’’ type of urolith
treated.

Eﬀectiveness for canine nephroliths

In 1996, Block and Adams

reported their initial experience with

ESWL in 5 dogs. Later, Adams and Senior

summarized their further

experience with nephroliths and ureteroliths in more than 30 dogs. Ongoing
treatment of many additional dogs, mostly with calcium oxalate, has made
Adams’ experience base the largest in the world. Using a Dornier HM-3
(water bath) lithotriptor, successful fragmentation is achieved in 90% of
dogs after one or two treatments

. One detailed report of canine

lithotripsy using a ‘‘dry’’ lithotriptor has been published

. Fragmentation

and reduction in size of two calcium oxalate ureteroliths and a nephrolith
were observed after ESWL (using a Lithostar II; Siemens Medical Systems,
Iselin, NJ) in a Golden Retriever dog. Three thousand to 4000 SWs at 16.6
to 17.8 kV were applied to each ureterolith, and an additional 2900 SWs at
16.3 kV were applied to the nephrolith to gain satisfactory initial
fragmentation. Fragments passed to clear the dog of uroliths completely
within 2 months

. We have summarized our initial experience with the

Modulith SL-20

and have now performed more than 30 treatments.

Nephroliths require 900 to 3900 SWs (average of about 2400 SWs) typically
at 16 to 18 kV for suﬃcient fragmentation in situ (

). Fragments begin

to move out of the renal pelvis within 24 hours but may take several weeks
to months to clear the upper urinary tract completely. By comparison, SW
doses for the ‘‘wet’’ Dornier HM-3 lithotriptor typically range from 750 to
1200 SWs at 13.5 to 15 kV, with a reported average SW dose of 900 SWs for
calcium oxalate nephroliths

. Retreatment is required in approximately

30% of cases with the Dornier HM-3

, whereas we have found an

approximately 50% retreatment rate, primarily for larger stones. Our
retreatment rate seems to be declining with continued improvement in
patient selection and technique.

Complications of extracorporeal shock wave lithotripsy for nephroliths

SW lithotripsy is inherently traumatic to kidney tissue. Possible renal

complications include renal swelling, renal or perirenal hemorrhage,
vasoconstriction, tubular damage, intrarenal scarring, and renal failure
(

[10–15]

. Long-term eﬀects of ESWL are still unclear in human

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

patients. In veterinary patients, transient hematuria is common after
lithotripsy

[7,8]

. Displacement of nephrolith fragments, ureterectasia,

retroperitoneal ﬂuid, and subcapsular hematoma formation were seen
sonographically in 14 dogs examined the day after lithotripsy, but no
change in echogenicity or architecture of the kidneys was detected

. In

our experience, fragment displacement into the ureter or renal calices is
common on posttreatment ultrasound imaging, whereas mild ureteral
dilation and perirenal ﬂuid are less common ﬁndings.

Minor transient changes in posttreatment creatinine measurements (\0.5

mg/dL) were described by Adams and Senior

in four dogs with

preexisting azotemia. In our experience, serum creatinine measurements
have remained in the normal range for 24 hours after treatment but have
increased from pretreatment measurements by 0.1 to 0.7 mg/dL in about
half of our treated dogs despite aggressive diuresis. The postlithotripsy
glomerular ﬁltration rate as determined by nuclear scintigraphy has
decreased from pretreatment measurements in seven of eight dogs assessed
between 7 and 60 days after treatment. Function may decline in a single
treated kidney or in both kidneys, supporting global hemodynamic eﬀects of
lithotripsy. The glomerular ﬁltration rate rebounded within several months
in some dogs when re-evaluated. Progressive renal failure has only been
observed in two of our treated dogs over the long term; both dogs had large
stone burdens and chronic pyelonephritis before treatment

SW damage is generally dependent on SW dose (power and number), and

although it is diﬃcult to compare energy applied by diﬀerent sources, the
higher SW doses required by the dry lithotriptor may more dramatically
aﬀect renal function in the short term. Methods to minimize renal injury are

Fig. 4. Example of nephrolith fragmentation. (A) Radiographic image before extracorporeal
shock wave lithotripsy treatment. (B) Radiographic image after application of 1300 and 2500
high-frequency shock waves to the left and right kidneys, respectively.

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

under investigation and may include changes in SW rate, modiﬁcations of
power setting, adjustment of lithotriptor energy delivery, or adjunct use of
renoprotective agents

[6,17–19]

Extrarenal complications include pain, bowel damage and diarrhea,

biochemical increases in amylase and lipase, and biochemical increases in
liver enzymes. Overt pancreatitis has been observed in veterinary (L.G.
Adams, DVM, PhD, personal communication, 2003) and human patients

[15,20]

Extracorporeal shock wave lithotripsy for ureteroliths

Lithotripsy fragmentation of ureteroliths is considered more diﬃcult than

that of nephroliths for several reasons. Clear imaging and accurate focusing
of ureteroliths may be more diﬃcult because of their small size and the
interference of various overlying tissues and is especially diﬃcult if
ultrasound is the primary imaging modality used. Careful positioning of
the ureterolith in the treatment focal zone is required to maximize SW
contact with ureterolith surfaces (

). Additionally, during lithotripsy,

Box 1. Possible complications of extracorporeal shock wave
lithotripsy

Immediate

Arrhythmia
Systemic hypertension
Hematuria
Perirenal hematoma
Pain

Early posttreatment

Hematuria
Renal hemorrhage and vascular injury
Pain
Ureteral obstruction
Decreased renal blood flow (RBF) and glomerular filtration rate
Hepatic enzyme elevation
Pancreatitis
Gastric or intestinal erosion, diarrhea

Delayed posttreatment

Systemic hypertension
Reduced renal function
Fibrosis and progressive chronic renal failure
Increased rate of stone recurrence

Data from Refs.

[3,9,15]

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

the ureterolith fragments may adhere to the ureteral mucosa and do not
immediately fall away from the original site, diminishing the eﬀectiveness of
subsequent SWs. Minimal movement and minimal reverberation of SWs
within the stone also limit the fragmentation eﬀect in the conﬁned space of
the ureter

[3,15]

. Higher SW doses and power settings are required; however,

SWs directed at ureteroliths are unlikely to be damaging to kidneys except
with extremely proximal ureteroliths. Factors limiting successful fragmen-
tation in human patients and leading to increased diversion to ureteroscopic
techniques have included larger stone size (>10–12 mm), distal (pelvic)
location

, degree of obstruction, and obesity

Using the dry lithotriptor, we have successfully treated ureteroliths in 10

dogs using an average of 2600 SWs at 14 to 19 kV. Only one ureterolith
lodged in the midureter in a small dog was insuﬃciently fragmented to pass
after initial treatment. By comparison, retreatment rates for ureteroliths were
greater than 50% (4 of 7 dogs required retreatment) using the Dornier HM-3
lithotriptor

. The primary complication of ureterolith fragmentation is

further ureteral obstruction. Fragmentation or movement of an ureterolith

Fig. 5. Example of ureterolith fragmentation. (A) Radiographic image during extracorporeal
shock wave lithotripsy treatment. (B) Radiographic image after application of 3500 high-
frequency shock waves to the distal ureterolith.

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

can create a more lodged stone, even if the ureterolith was nonobstructive
initially. In our experience, ESWL treatment of ureteroliths can be more
painful than treatment of nephroliths alone. If the ureterolith is reasonably
separated from the kidney, renal or pancreatic damage is unlikely.

Extracorporeal shock wave lithotripsy in feline patients

Feline uroliths seem to be more diﬃcult to fragment with ESWL than

canine uroliths, and feline kidneys may be particularly sensitive to ESWL.
Using a Dornier HM-3 lithotriptor, Adams and Senior

found that

ureteroliths could be fragmented successfully in only one of ﬁve cats and
that fragmentation of nephroliths was incomplete as well. In addition,
transient or permanent worsening of renal function occurred in several cats.
Gonzales et al

recently investigated the eﬀects of dry lithotripsy on the

kidneys in four healthy cats. Using a Dornier MFL-5000 lithotriptor, the
investigators found no signiﬁcant eﬀects on ultrasonographic appearance of
the kidneys at 24 hours and 14 days after treatment (2000 SWs at 24 kV).
Additionally, there was no diﬀerence in pretreatment and 14-day posttreat-
ment glomerular ﬁltration rate measurements in these cats

. Although

these results are promising, this SW dosage may not be adequate to
fragment feline uroliths suﬃciently and diseased feline kidneys may respond
diﬀerently to the ESWL insult than healthy kidneys. Adams et al

studied the in vitro fragmentation of intact calcium oxalate uroliths
retrieved from dogs and cats and submitted to the Minnesota Urolith
Center. Using a research electrohydraulic lithotriptor that simulates the
function of the Dornier HM-3, breakage of the stones was evaluated using
digital image size. In this study, signiﬁcantly less breakage was observed in
feline stones than in canine uroliths after the same SW dose (100 SWs at 20
kV)

, conﬁrming clinical evidence that feline uroliths are simply ‘‘tougher

nuts to crack.’’

Currently, we consider treating obstructive ureteroliths in cats with

informed owner consent. In one cat with multiple ureteroliths, we
documented improved azotemia after lithotripsy despite little change in
the appearance of the ureteroliths. In another cat, successful fragmentation
of a partially obstructive ureterolith was obtained after two lithotripsy
treatments, both applying more than 3500 SWs at 18 to 19 kV. Increased
obstruction, however, was observed between treatments. Because of the
diﬃculty in fragmenting feline uroliths, we are hesitant to treat feline
nephroliths at this time unless they are associated with signiﬁcant infection,
destruction of the kidney, or obstruction of urine ﬂow.

Limitations of extracorporeal shock wave lithotripsy

The most common reason why lithotripsy is not recommended for

speciﬁc uroliths is that intervention is unnecessary. Many canine and feline

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

nephroliths remain static in size and clinically silent for years. For dogs with
struvite, calcium oxalate, and urate stones in which intervention is indicated,
there are few contraindications to lithotripsy. Absolute contraindications to
lithotripsy in human patients include pregnancy and untreated coagulop-
athy

. The eﬀectiveness of ESWL is also limited with extremely large

stones (>20 mm in diameter)

. Relative contraindications to ESWL in

veterinary patients include systemic hypertension, overt hyperadrenocorti-
cism, or conditions that would compromise the patient’s ability to handle
either general anesthesia or posttreatment diuresis.

Although persistent infection is a common indication for intervention

with nephroliths, active pyelonephritis should be appropriately managed
before ESWL. Periprocedural antimicrobials are indicated in patients with
a history of persistent or recurrent UTI. We have seen one patient with
negative pretreatment urine cultures exhibit bacteriuria after fragmentation
of nephroliths.

The major limitations of ESWL are the structural limitations on

locations that can be treated with this method. For EWSL to be eﬀective,
the focal zone must be positioned in an anatomic window in which SWs can
be transmitted easily to the stone through a continuous ﬂuid or soft tissue
medium and without interference from bony structures. This feature makes
the procedure impractical for urethral stones and diﬃcult for distal
ureteroliths in extremely small animals. Although rarely encountered in
small animals, patient positioning and depth of penetration of SWs is
limited in patients weighing greater than 150 kg.

Another limitation of lithotripsy is the possibility that the procedure may

not completely rid the kidney of stone material, especially with larger stones.
Small (\2 mm) remaining fragments are unlikely to be clinically signiﬁcant

[3,7,15]

unless they continue to serve as an infection nidus. It is also diﬃcult

to analyze stone material unless a fragment can be collected by spontaneous
voiding or by voiding hydropropulsion or other nonsurgical methods.
ESWL is also of limited eﬀectiveness with certain ‘‘hard’’ stone types,
including cystine and calcium oxalate dihydrate uroliths

[3,25]

Extracorporeal shock wave lithotripsy for cystouroliths

ESWL is not widely recommended for treatment of bladder stones,

because mobile bladder stones are readily able to move out of the SW path
within the bladder lumen. This movement limits the repeated SW eﬀect on
cystoliths and may result in larger fragments than desired. Extracorporeal
lithotripsy can, however, be used to reduce the size of cystoliths for either
dissolution or hydropropulsion. Successful fragmentation of bladder stones
using a water bath lithotriptor has been reported in a dog

. Adams and

Senior

found fragmentation of bladder stones adequate in only two of

ﬁve dogs using a similar method. We have been pleased with the ability of
the dry lithotriptor to fragment bladder stones for suﬃcient passage in

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

several female dogs and one cat but have avoided this treatment in male
dogs because of the increased risk of urethral obstruction. We currently
consider treatment of larger or multiple calcium-containing bladder stones
in patients that have had multiple cystotomies, are poor surgical candidates,
or are being treated concurrently for upper urinary tract stones. We also
consider ESWL fragmentation of large or multiple struvite uroliths in
female dogs in an attempt to hasten medical dissolution. Intracorporeal
methods are preferred for optimal fragmentation of cystoliths, however.

Intracorporeal lithotripsy

Mechanical, pneumatic, electrohydraulic, and laser modalities have

expanded the abilities of intracorporeal lithotripsy over the last 50 years

[3,27,28]

. Initially used as direct contact methods for accessible bladder

stones, progressively smaller sizes of instruments have allowed percutaneous
and endoscopic access to pelvic, ureteral, and lower tract calculi. EHL has
been described

[3,27,29]

and remains the most economic method for

veterinary application. Instrumentation required includes a cystoscopic
system, EHL spark generator, and disposable electrode probes. Although
a variety of probe sizes are available, most require at least an 11-French
cystoscope sheath, with 17- to 20-French sheaths preferred

. Larger

sheaths are essential for adequate ﬂushing and retrieval of the irregular
urolith fragments caused by EHL. An experienced operator and generous
working room in the urinary bladder are required as well, because the EHL
SWs can be damaging if discharged too close to the bladder wall. Hemorrhage
and mucosal damage may ensue, and perforation can occur if the probe is
discharged at high-power settings when in contact with urothelial surfaces.
This potential damage at high-power settings also precludes the eﬀective use of
EHL probes in locations like the ureter or urethra

Laser methods are rapidly supplanting other intracorporeal methods and

likely represent the state of the art for future treatment of urethroliths and
ureteroliths in larger dogs. As early as 1968, a long-pulsed ruby lithotriptor
was developed that could fragment calculi; however, limitations of this and
other early lasers were surrounding tissue damage

. More recently, the

pulsed neodymium:yttrium-aluminum-garnet (Nd:YAG), pulsed-dye, alex-
andrite, and holmium:yttrium-aluminum-garnet (Ho:YAG) lithotriptors
have been safely introduced into clinical use

[27,30]

. Pulsed-dye lasers

create urolith fragmentation by a combination of eﬀects at the surface of the
urolith. Energy plasma is generated by the photoacoustic eﬀects of the laser
and is followed by an expanding cavitation bubble. Collapse of the
cavitation bubble is thought to generate the SWs that ultimately fragment
the stone

. Pulsed-dye lasers have been used successfully for equine

urolithiasis

but have been labor-intensive to maintain

The Ho:YAG laser uses the active medium of holmium, a rare earth

element, and the YAG crystal to operate at a wavelength of 2100 nm,

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

creating photothermal energy that can be delivered through small ﬂexible
ﬁbers. Pulse durations range from 250 to 350 microseconds, pulse energies
vary from 200 to 4000 mJ, frequencies vary from 5 to 45 Hz, and power
outputs vary from 31 to 80 W. The ﬂexible laser ﬁbers range from 200 to 100
lm in size, with the smaller sizes providing the most eﬀective stone ablation

. Direct contact with the urolith in liquid media allows vaporization of

water molecules to create the cavitation bubble

. Laser lithotripsy with

the Ho:YAG laser (360-lm ﬁber) has been successful in fragmenting canine
uroliths in vitro without damaging the optical ﬁbers

. Clinically, Halland

et al

described successful obliteration of urethroliths in three goats and

two pot-bellied pigs using a 220-lm Ho:YAG ﬁber directed through an 8-
French (2.7-mm) ureteroscopic instrument. A smaller 5.5-French 70-cm
endoscope was found to be too fragile to navigate the urethra. The Ho:YAG
laser was also useful in reducing the size of calcium carbonate cystoliths in
horses to facilitate removal by other nonsurgical methods

. Davidson et al

recently described the application of Ho:YAG laser energy to surgically

implanted urethroliths placed at the base of the os penis of healthy dogs.
Although most urethroliths were fragmented small enough to be ﬂushed out
of the urethra, incomplete fragmentation, retropulsion of fragments into the
urinary bladder, and urethral mucosal damage are potential limitations of
this technique in clinically aﬀected dogs

. The eﬀectiveness of laser

lithotripsy may vary with diﬀerent species or stone types, but it seems to be
the most promising device for canine applications

. Other urologic

applications of laser lithotripsy include structure incision, prostate resection,
and ablation of superﬁcial bladder tumors

. Although laser lithotripsy is

considered safer than EHL in conﬁned spaces, such as the ureter and
urethra, repeated pulses on the urothelium can cause perforation

http://www.vet.utk.edu/clinical/sacs/lithotripsy.

The expense of laser generators (an investment of $80,000–$100,000) has

been prohibitive for veterinary medicine in the past; however, leasing
arrangements have been used by some to avoid the capital expense of the
system. Additionally, the Ho:YAG laser can be applied in procedures across
specialties (eg, surgery, ophthalmology) and may be a feasible equipment
purchase for large specialty practices or veterinary teaching hospitals.

Appendix A: selected sites for small animal lithotripsy

Purdue University Veterinary Teaching Hospital
1248 Lynn Hall
West Lafayette, IN 47907–1248(765)-494-1107
Contact: Dr Larry Adams
Extracorporeal show wave lithotripsy (ESWL), electrohydraulic shock wave
lithotripsy (EHL), and holmium:yttrium-aluminum-garnet (Ho:YAG) laser

The University of Tennessee College of Veterinary Medicine
Veterinary Teaching Hospital

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I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

Knoxville, TN 37996–4544(865) 974-8387
Contact: Mot Daughtridge or Dr India Lane
Information available at:

shtml

ESWL

Foster Hospital for Small Animals
Tufts University
200 Westboro Road
North Grafton, MA 01536(508) 839-5395
Contact: Dr Mary Ann Labato or Dr Linda Ross
ESWL

Oklahoma State University
Boren Veterinary Teaching Hospital
Stillwater, OK 74078(405) 744-7000
Contact: Dr Ellen Davidson
Ho:YAG Laser

References

[1] Chaussy C, Brendel W, Schmiedt E. Extracorporeally induced destruction of kidney stones

by shock waves. Lancet 1980;2:1265–8.

[2] Chaussy C, Schmidt E, Jocham D, Brendel W, Forrsman B, Walther V. First clinical

experience with extracorporeally induced destruction of kidney stones by shock waves.
J Urol 1982;127:417–20.

[3] Adams LG, Senior DF. Electrohydraulic and extracorporeal shock-wave lithotripsy. Vet

Clin North Am Small Anim Pract 1999;29:293–302.

[4] Preminger GM. Shock wave physics. Am J Kidney Dis 1991;17:431–5.
[5] Chow GK, Streem SB. Extracorporeal lithotripsy: update on technology. Urol Clin North

Am 2000;27(2):315–22.

[6] Auge BK, Preminger GM. Update on shock wave lithotripsy technology. Curr Opin Urol

2002;12:287–90.

[7] Block G, Adams LG, et al. The use of extracorporeal shock wave lithotripsy for treatment

of spontaneous nephrolithiasis and ureterolithiasis in dogs. J Am Vet Med Assoc 1996;208:
531–6.

[8] Bailey G, Burk RL. Dry extracorporeal shock wave lithotripsy for treatment of

ureterolithiasis and nephrolithiasis in dog. J Am Vet Med Assoc 1995;207:592–5.

[9] Lane IF. Dry extracorporeal lithotripsy in small animals. In: Proceedings of the 21st Annual

Forum of the American College of Veterinary Internal Medicine, Charlotte, NC, 2003. p.
775–6.

[10] Delius M, Enders G, Xuan ZR, Liebich HG, Brendel W, et al. Biological eﬀects of shock

waves: kidney damage by shock waves in dogs—dose dependence. Ultrasound Med Biol
1988;14:117–22.

[11] Karlsen SJ, Smevik B, Strenstrom J, Berg KJ. Acute physiological changes in canine

kidneys following exposure to extracorporeal shock waves. J Urol 1990;143:1280–3.

[12] Rassweiler J, Kohrmann KU, Back W, et al. Experimental basis of shock wave-induced

renal trauma in the model of the canine kidney. World J Urol 1993;11:43–53.

[13] Newman R, Hackett R, Senior D, et al. Pathologic eﬀects of ESWL on canine renal tissue.

Urology 1987;29:194–200.

1024

I.F. Lane / Vet Clin Small Anim 34 (2004) 1011–1025

[14] Willis LR, Evan AP, Connors BA, Blomgren P, Fineberg NS, Lingeman JE. Relationship

between kidney size, renal injury and renal impairment induced by shock wave lithotripsy.
J Am Soc Nephrol 1999;10:1753–62.

[15] Lingeman JE, Lifshitz DA, Evan AP. Surgical management of urinary lithiasis. In: Walsh

PC, editor. Campbell’s urology. 8th edition. Philadelphia: WB Saunders; 2002. p. 3361–452.

[16] Siems JJ, Adams LG, et al. Ultrasound ﬁndings in 14 dogs following extracorporeal shock-

wave lithotripsy for treatment of nephrolithiasis [abstract]. In: Proceedings of the
American College of Veterinary Radiology, Chicago, IL, 1999. p. 11.

[17] Munver R, Delvecchio FC, Kuo RL, Brown SA, Zhong P, Preminger GM. In vivo

assessment of free radical activity during shock wave lithotripsy using a microdialysis
system: the renoprotective action of allopurinol. J Urol 2002;167:327–34.

[18] Paterson RF, Kuo RL, Lingeman JE. The eﬀect of rate of shock wave delivery on the

eﬃciency of lithotripsy. Curr Opin Urol 2002;12(4):291–5.

[19] Sokolov DL, Bailey MR, Crum LA. Use of a dual-pulse lithotripter to generate a localized

and intensiﬁed cavitation ﬁeld. J Acoust Soc Am 2001;110:1685–95.

[20] Abe H, Nisimura T, Osawa S, Miura T, Oka F. Acute pancreatitis caused by extracorporeal

shock wave lithotripsy for bilateral renal pelvic calculi. Int J Urol 2000;7(2):65–8.

[21] Shiroyanagi Y, Yagisawa T, Nanri M, Kobayashi C, Toma H. Factors associated with

failure of extracorporeal shock-wave lithotripsy for ureteral stones using Dornier
lithotripter U/50. Int J Urol 2002;9(6):304–7.

[22] Delakas D, Karyotis I, Daskalopoulos G, Lianos E, Mavromanolakis E. Independent

predictors of failure of shockwave lithotripsy for ureteral stones employing a second-
generation lithotripter. J Endourol 2003;17(4):201–5.

[23] Gonzales A, Labato M, Solano M, Ross L. Evaluation of the safety of extracorporeal

shock-wave lithotripsy in cats [abstract]. In: Proceedings of the 20th American College of
Veterinary Internal Medicine Forum, Dallas, TX, 2002. p. 810.

[24] Adams LG, Williams JC Jr, McAteer JA, et al. In vitro evaluation of canine and feline

urolith fragility by shock wave lithotripsy [abstract]. J Vet Intern Med 2003;17(3):406.

[25] Williams JC, Chee Saw K, Paterson RF, et al. Variability of renal stone fragility in shock

wave lithotripsy. Urology 2003;61(6):1092–6.

[26] Loske AM, Prieto FE, Lopez JA. Primer tratamiento de litotripsia extracorporal en un

perro usando ungenerador de ondas de choque experimental hecho en Mexico. Vet Mex
1996;27:41–8.

[27] Zheng W, Denstedt JD. Intracorporeal lithotripsy: update on technology. Urol Clin North

Am 2000;27(2):301–13.

[28] Senior DF. Electrohydraulic shock-wave lithotripsy in experimental canine struvite

bladder stone disease. Vet Surg 1984;13:143–8.

[29] Eustace RA, Hunt JM. Electrohydraulic shock-wave lithotripsy for the treatment of cystic

calculus in two geldings. Equine Vet J 1988;20(3):221–3.

[30] Bartels KE. Lasers in veterinary medicine—where have we been and where are we going?

Vet Clin North Am Small Anim Pract 2002;32(3):495–515.

[31] Howard RD, Pleasant RS, May KA. Pulsed dye laser lithotripsy for treatment of

urolithiasis in two geldings. J Am Vet Med Assoc 1998;212:1600–3.

[32] Wynn VM, Davidson EB, Higbee RG, Ritchey JW, Ridgway TD, Bartels KE, et al. In

vitro eﬀects of pulsed holmium laser energy on canine uroliths and porcine cadaveric
urethra. Laser Surg Med 2003;3:243–6.

[33] Halland SK, House J, George LW. Urethroscopy and laser lithotripsy for the diagnosis

and treatment of obstructive urolithaisis in goats and pot-bellied pigs. J Am Vet Med
Assoc 2002;220(12):1831–4.

[34] Judy CE, Galuppo LD. Endoscopic-assisted disruption of urinary calculi using

a holmium:YAG laser in standing horses. Vet Surg 2002;31(3):245–50.

[35] Davidson EB, Ritchey JW, Higbee RD, et al. Laser lithotripsy for treatment of canine

uroliths. Vet Surg 2004;33(1):56–61.

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Urine culture as a test for cure:

why, when, and how?

Jody P. Lulich, DVM, PhD*,

Carl A. Osborne, DVM, PhD

Minnesota Urolith Center, College of Veterinary Medicine, University of Minnesota,

1352 Boyd Avenue, St. Paul, MN 55108, USA

Case scenario

Consider the following case scenario. A colleague in private practice

called the urology service for advice about a 9-year-old-spayed female
Golden Retriever with persistent pollakiuria and gross hematuria of 2
months’ duration. Urinalysis revealed numerous red blood cells, white
blood cells, proteinuria, and spherical structures thought to be bacterial
cocci. A diagnosis of bacterial urinary tract infection (UTI) was made,
and therapy with orally administered amoxicillin with clavulanic acid was
prescribed. After 2 weeks of therapy, the owners indicated that clinical
signs were still present but less severe. Urinalysis revealed persistence of
numerous red blood cells and white blood cells. Based on the assumption
that uropathogenic bacteria causing these abnormalities were resistant to
amoxicillin with clavulanic acid, oral administration of trimethoprim/
sulfadiazine was provided. In this clinical setting, what is the likelihood
that poor response to therapy was caused by antimicrobial resistance by
uropathogenic bacteria? Furthermore, what is the likelihood that use of
trimethoprim/sulfadiazine will be associated with eradication of pollakiu-
ria and hematuria?

In considering the answers to these questions, consider the following

observations. In two retrospective clinical studies, it was estimated that the
hospital proportional morbidity rate of UTI in dogs was 10% to 14%

. In contrast, in another retrospective clinical study, the hospital

proportional morbidity rate for persistent or recurrent UTIs in dogs was
only 0.3%

lulic001@maroon.tc.umn.edu

. In a retrospective clinical study of 100 dogs with persistent

* Corresponding author.
E-mail address:

(J.P. Lulich).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.005

Vet Clin Small Anim

34 (2004) 1027–1041

or recurrent infections, only 30% of 374 bacterial isolates were resistant to
achievable serum concentrations of commonly prescribed oral antibiotics
(ie, trimethoprim-sulfonamides, amoxicillin, amoxicillin with clavulanic
acid, cephalexin, tetracycline)

. When considered in the context of the

fact that these antimicrobics usually achieve higher concentrations in urine
than in serum, 30% is likely to represent an overestimation of the true
prevalence of resistant microbes as a cause for persistent or recurrent
UTIs.

In the context of the Golden Retriever described in the clinical scenario,

which of the following two assumptions is the most likely explanation for
persistent clinical signs of lower urinary tract disease?

1. Pollakiuria, hematuria, and inﬂammation detected by urinalysis were

caused by uropathogenic bacteria that were resistant to amoxicillin with
clavulanic acid therapy.

2. Pollakiuria, hematuria, and inﬂammation detected by urinalysis were

caused by a noninfectious disease process that would not respond to
amoxicillin with clavulanic acid therapy.

In contemplating the answer to this question, consider the following.

Clinical signs persisted for weeks despite therapy with trimethoprim/sulfadi-
azine. Because of the presumption that the dog had UTI caused by resistant
bacteria, it was subsequently treated with two additional antimicrobial drugs
without success. At that point, radiography and biopsy of the urinary bladder
were performed. The results revealed an inﬁltrating transitional cell carcinoma
of the urinary bladder. It is apparent that treatment failure in this instance was
not a result of the use of ineﬀective antimicrobial drugs to treat resistant
uropathogens. Rather, it was associated with ineﬀective use of antimicrobial
drugs to treat a noninfectious disorder.

What is the point? In addition to bacterial infections, many diverse

noninfectious disease processes, including neoplasia and urolithiasis, result
in inﬂammatory lesions of the urinary tract characterized by exudation of
red blood cells, white blood cells, and protein into urine. Resultant
hematuria, pyuria, and proteinuria suggest inﬂammatory urinary tract
disease. As with the patient described here, diagnosis of bacterial UTI
solely on the basis of urinalysis and detection of inﬂammatory cells in urine
sediment results in overdiagnosis. Therefore, it is essential to distinguish
between inﬂammation and infection related to urinary tract disease.
Although detection of bacteria in fresh urine sediment should prompt
consideration of UTI, it should be veriﬁed by urine culture. Nonbacterial
‘‘look-alikes’’ in urine sediment are often confused with bacteria.

In retrospect, how would you modify management of the urinary tract

disorder in this female Golden Retriever? Won’t you agree that evaluation
of in vitro cultures of urine for bacterial pathogens was warranted before the
initiation of therapy?

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What are diagnostic urine cultures?

Quantitative urine culture before initiation of antimicrobial therapy is

considered to be the gold standard for diagnosis of bacterial UTIs. If the
patient is currently being treated with an antimicrobic, it should be
discontinued for approximately 3 to 5 days before diagnostic urine culture
to minimize inhibition of in vivo and in vitro bacterial growth. In addition
to facilitating diﬀerentiation of harmless bacterial contaminants from
bacterial pathogens, accurate identiﬁcation of speciﬁc bacterial species
aids in selection of antimicrobial drugs. It also facilitates diﬀerentiation
of recurrent UTIs caused by relapses from recurrent UTIs caused by
reinfections. UTIs caused by relapses cannot be distinguished from re-
current UTIs caused by reinfections without comparison of pretreatment
bacterial culture results with follow-up culture results. The point to be
emphasized is that failure to perform bacterial urine cultures or failure to
interpret results of urine cultures correctly may lead not only to diagnostic
errors but to therapeutic failures as well.

What are therapeutic urine cultures?

In addition to diagnostic culture of urine before treatment of the patient

with antimicrobics, culture of urine at strategic times during antimicrobial
therapy (so-called ‘‘therapeutic urine cultures’’) is an eﬀective method of
assessing therapy. If patients are not responding to treatment, therapeutic
cultures are essential for determining why there is a lack of response (

When should therapeutic urine cultures be considered?

The need to obtain cultures during therapy depends on severity of

disease, underlying reasons for antimicrobic use, risks associated with
antimicrobic use, and health of the patient. The beneﬁts of therapeutic urine
cultures are listed in

For patients with a high risk of morbidity and mortality (eg, prostatitis,

pyelonephritis, immunosuppression, urinary tract obstruction), evaluation
of a urine culture and urinalysis 3 to 5 days after initiating therapy allows
veriﬁcation of antibiotic eﬀectiveness before development of irreversible
organ damage or systemic spread of disease. The same strategy should be
considered when prescribing antimicrobics with a high risk of toxicity.
Suggested times to culture urine to diagnose and monitor persistent UTIs
are listed in

In general, we recommend culture of urine 3 to 5 days after initiation of

antimicrobic therapy because it facilitates timely assessment of in vivo
eﬀectiveness of treatment. Initial response to therapy is considered to be
eﬀective only if in vitro culture of properly collected urine samples does not

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CLINICAL SIGNS PERSIST DURING ANTIMICROBIC THERAPY

No Bacterial Growth

Restore host defenses
Antimicrobic eradication of infection or
Delay/postpone antimicrobics

Bacteria are susceptible to

current antimicrobics

Bacteria are NOT susceptible to

current antimicrobics

INADEQUATE ANTIMICROBIC DELIVERY

ANTIMICROBIC RESISTANCE

Select active drug
Administer sufficient dose
Improve client compliance and patient acceptance
Promote intestinal absorption
Minimize drug inactivation
Expose/eradicate infection nidus

Restore host defenses
Mono/multidrug therapy or
Delay/postpone antimicrobics

New Bacterial Isolate

Original Bacterial Isolate

Investigate other causes:

Structural abnormalities
Functional abnormalities
Behavioral changes

NONBACTERIAL DISEASE

SUPERINFECTION

Antimicrobial Susceptibility Results

Aerobic Culture Results

Fig. 1. Algorithm for diagnosis and management of antimicrobic failure in patients with
urinary tract infection.

Box 1. Benefits of therapeutic urine cultures

1. Timely test of antimicrobic efficacy
2. Verification of proper antimicrobic administration
3. Early detection of bacterial resistance to antimicrobics
4. Timely detection of persistent infections
5. Provision of justification for early discontinuation of potentially

toxic antimicrobics

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result in growth of bacteria. Even though there may be viable bacterial
pathogens in surrounding tissues, the urine containing an appropriate
concentration of antimicrobic should be sterile. Treatment is ineﬀective and
relapse occurs if the bacterial colony count has only been reduced (eg, from
10

to 10

). Therefore, one cannot rely on examination of urine sediment to

detect elimination of bacteria by therapy, because persistence of reduced
numbers of bacteria may fall below the limit of detection by light
microscopic evaluation of urine sediment. Although urine contains no
viable microbes, hematuria, pyuria, and proteinuria associated with
compensatory inﬂammation are likely to be detected by urinalysis 3 to 5
days after initiating eﬀective antimicrobial therapy. These inﬂammatory-
associated abnormalities may be of lesser magnitude compared with
pretreatment ﬁndings, however. Anytime clinical signs persist after a suitable
period of antimicrobic therapy, therapeutic urine cultures should also be
repeated.

For complicated UTIs, consider evaluation of a urine culture and urinalysis

3 to 5 days (or sooner if necessary) before scheduled discontinuation of
therapy, especially if prophylactic antibiotics are to be subsequently used to
prevent frequent reinfection. Therapy may be discontinued if urine is sterile
and urine sediment is normal. If culture results indicate persistent infection,
however, re-evaluation of therapy is essential. In this situation, initiation of
prophylactic low-dose antimicrobial therapy is contraindicated.

How should urine samples be collected for bacterial culture?

We prefer to collect urine samples for bacterial culture by cystocentesis to

avoid iatrogenic UTI and to eliminate problems of diﬀerentiating contam-
inants from pathogens. Detection of bacteria, even in low numbers, in urine

Box 2. Suggested times to culture urine to diagnose
and monitor persistent urinary tract infections

I. Diagnostic

Before administration of therapy

II. Therapeutic

3 to 5 days after initiating therapy
Any time clinical signs or laboratory abnormalities recur

during therapy

Before discontinuing therapy

III. Surveillance

7 to 14 days after stopping therapy
1 to 2 months after stopping therapy
3 to 6 months after stopping therapy
Any time clinical signs recur

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aseptically collected by cystocentesis is indicative of UTI. False-positive
results may occur if the needle penetrates a loop of intestine during
cystocentesis or if the sample is contaminated during transfer to culture
media, however. For this reason, quantitative urine culture is routinely
recommended, even for samples collected by cystocentesis. In addition, urine
culture results should be interpreted in the context of other clinical ﬁndings.

How should urine samples be preserved before culture?

Because urine may be a good culture medium at room temperature

(bacterial counts may double every 20–45 minutes), it should be cultured
within 15 to 30 minutes from the time of collection. Another indication for
culture of fresh urine samples is that destruction of some fastidious bacteria
may occur within an hour of collection. If culture of freshly collected urine
samples is not possible for any reason, samples should be kept in a sealed
sterile container and immediately refrigerated after collection. Refrigerated
samples may be stored for 6 to 12 hours without signiﬁcant additional
growth of bacteria; nevertheless, it is emphasized that fastidious organisms
may be killed in the urine environment if refrigeration storage time is
prolonged. Freezing urine samples may also destroy bacteria.

How do therapeutic cultures help to detect and manage antimicrobic failures?

Failure to respond to antimicrobic drug therapy results from a variety of

causes. Potential causes for poor response to antimicrobic therapy are listed
in

Box 3

Empirically switching the antimicrobic or adding additional antimicro-

bics based on the presumption that a resistant uropathogen is present may
be eﬃcient but is often ineﬀective. To minimize expense and iatrogenic
morbidity associated with ineﬀective antimicrobial therapy, appropriate
diagnostic tests to determine the cause of failure should be performed ﬁrst.
In this context, we recommend evaluation of urine culture and antimicrobic
susceptibility results before altering antimicrobial therapy (see

Although the following sections of this article focus on therapeutic failures
related to ineﬀective use of antimicrobial drugs and use of ineﬀective
antimicrobial drugs, recall that establishment of infections is related to
impaired host defense mechanisms in addition to virulence of bacterial
pathogens. In a retrospective study, underlying disorders capable of
breaching normal host defenses were identiﬁed in 71% of dogs with
persistent and recurrent UTIs

. When underlying causes were corrected,

control of infection improved in almost half of the dogs. These ﬁndings
indicate that identiﬁcation and correction of underlying defects in host
defenses also represent an important strategy for success.

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Culture Result

¼ No Bacterial Growth

Negative therapeutic culture results during antibiotic administration are

consistent with successful eradication of infection. Therefore, persistence of
clinical signs should prompt further evaluation of the patient (eg, transrectal
palpation of the urethra, radiographic and ultrasonographic imaging,
cystoscopy) for concomitant nonbacterial disorders as causes for a UTI.

Box 3. Potential causes for poor response to antimicrobic
therapy

1. Administration of antimicrobics to correct noninfectious

disorders causing signs similar to those associated with
urinary tract infection (UTI)

2. Inadequate antimicrobic delivery

Poor client compliance

Unable to administer drug
Unwilling to adhere to dosage schedules
Premature withdrawal of drug as clinical signs subside
Discontinuation of drug because of expense

Poor patient acceptance

Unwilling to swallow drug
Vomiting after drug administration
Unable to tolerate side effects of drugs

Ineffective drug

Expired drug
Inactivated drug
Ineffective dose
Diuresis reducing urine drug concentration

Impaired drug transport

Decreased intestinal absorption
Altered perfusion of infected tissue
Microbes sequestered in inaccessible site
Decreased glomerular filtration reducing urinary

excretion

3. Microbes resistant to antimicrobic

Intrinsic resistance
Acquired resistance

4. Superinfection

Failure or inability to recognize and eliminate or control

predisposing cause for UTIIatrogenic reinfection caused by
catheters or surgical techniques

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As illustrated in the case scenario at the beginning of this discussion, dysuria
and hematuria associated with lower urinary tract neoplasia should not be
expected to resolve in association with antimicrobial therapy

False-negative cultures can result from iatrogenic bactericidal events

during collection, storage, or transport of urine samples to diagnostic
facilities. Therefore, appropriate sample handling should be maintained
when samples are shipped for processing.

Culture Result

¼ Isolation of the Same Bacterial Species Susceptible

to the Currently Administered Antimicrobic

Culture results that remain unchanged (ie, the same bacteria exhibiting in

vitro susceptibility to the current antimicrobic drug) during appropriate
antimicrobic therapy indicate that antimicrobics are not reaching the site of
infection. Incomplete or infrequent administration of antimicrobics and
unwillingness of patients to accept medications are common causes for
inadequate tissue delivery of apparently eﬀective medicine. In a study
conducted in conjunction with the American Animal Hospital Association,
only 48% of pet owners were compliant in administering heartworm
preventative

. We can expect a similar lack in compliance during

administration of antimicrobic drugs, especially as clinical signs wane. To
improve client compliance, consider medications that are easy to administer
and readily accepted by patients and administration regimens that can be
easily incorporated into owners’ schedules.

In some situations, antimicrobic administration is suﬃcient but intestinal

absorption is impaired. For example, tetracyclines have been recommended
as an eﬀective treatment for Pseudomonas UTI with a reasonable assurance
of success

. To facilitate oral administration, some owners package

medication in appetizing malleable foods, such as cheese. However, when
human patients were administered demeclocycline with 110 g of cottage
cheese, serum concentrations were 60% to 80% lower than when demeclo-
cycline was administered without food

. Tetracyclines, not including

doxycycline and minocycline, form relatively insoluble chelates with divalent
and trivalent metals, such as calcium, aluminum, magnesium, and iron.
Antacids containing aluminum, magnesium, and calcium also adversely
aﬀect absorption of tetracyclines and possibly other antimicrobics as well.
Therefore, in addition to evaluating whether or not medications are being
administered at the prescribed dosage and time intervals, the method of oral
administration should be considered as well.

Some diseases may also increase the risk of poor therapeutic response by

impairing adequate delivery of otherwise eﬀective antimicrobics to sites of
infection. For example, eradication of bacteria embedded inside uroliths or
bacteria walled oﬀ within abscesses is dependent on passive diﬀusion of an
adequate quantity of antimicrobic into site(s) of infection. Because these

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locations lack an intrinsic blood supply to ensure delivery of an adequate
quantity of antimicrobic, however, bacterial cure is unpredictable. There-
fore, in these situations, surgical drainage or eradication of the nidus of
infection remains an important component of successful treatment.

Recognition of recurrent UTIs associated with bacterial infections

inaccessible to antimicrobics that otherwise would eradicate them warrants
further consideration. Although viable bacteria remain sequestered in sites
inaccessible to bactericidal concentrations of antimicrobics, cultures of
samples obtained by cystocentesis during antimicrobic administration are
likely to be negative if the drug is eliminated in a high concentration in
urine. Urine cultures are likely to be positive after an interruption in
antimicrobic therapy, however. Consequently, these types of infections are
commonly diagnosed as relapses of a persisting uropathogen.

Inadequate antimicrobic delivery may also be associated with adminis-

tration of outdated or inactivated medications. We have experienced
situations where poor response to treatment of bacterial UTI may have
been associated with owners who removed tablets containing amoxicillin
with clavulanic acid from their protective foil covering weeks in advance to
facilitate administration. Clavulanic acid is susceptible to inactivation once
exposed to moisture. Similarly, inappropriately prescribing or administering
doses lower than recommended may reduce antimicrobic eﬀectiveness,
especially in polyuric patients.

Culture Result

¼ Isolation of the Same Bacterial Species That Is

Not Susceptible to the Currently Administered

Antimicrobic

Emerging antimicrobial resistance should be suspected when results of

therapeutic cultures reveal persistence of the original organism that is no
longer susceptible to antimicrobics that were previously eﬀective. Resistant
bacteria ﬂourish during ineﬀective antibiotic use because they have
a selective advantage over the remaining susceptible bacterial population.
Therefore, once resistance is detected, current antimicrobic therapy should
be discontinued. Selection of replacement antimicrobics should be based on
susceptibility results. To ensure that a suﬃcient concentration of drug is
maintained at the site of infection, administer medications at the higher end
of the dosage range (particularly when administering concentration-de-
pendent antimicrobics, such as ﬂuoroquinolones) and with increased
frequency (particularly when administering time-dependent antimicrobics,
such as potentiated penicillins).

Simultaneous administration of two or more antimicrobial agents is

based on the premise that antibacterial killing activity is enhanced, thereby
minimizing the development of antimicrobic resistance. Only a few

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antimicrobic combinations have been evaluated for treating UTIs in dogs
and cats, however. For example, in dogs, greater than 94% of Staphylococcus
intermedius

infections demonstrated in vitro susceptibility to the synergistic

eﬀects of amoxicillin with clavulanic acid, whereas only 20% were susceptible
to ampicillin without clavulanic acid

. Compared with other bacterial

species, Pseudomonas spp demonstrate a high frequency of resistance to
many antimicrobics in dogs

and human beings

. Because of the ability

of bacteria to adapt rapidly by a variety of pathways to avoid antimicrobic
killing, synergistic antimicrobic combinations have been recommended in
human beings

. Although antipseudomonal penicillins (ticarcillin and

piperacillin) and aminoglycosides have been historically prescribed, sub-
stitutions of newer antimicrobics (ie, third-generation cephalosporins or
ﬂuoroquinolones for antipseudomonal penicillins or aminoglycosides) may
be as eﬀective. In human beings, eﬃcacy of combination antipseudomonal
therapy has been reﬂected in improved survival rates in patients with
septicemia and neutropenia

. Based on these ﬁndings, we assume that

appropriate combination therapy for similar situations would eﬀectively
eradicate resistant bacteria and reduce emergence of resistant urinary tract
pathogens in dogs and cats. Nevertheless, there are also potential negative
aspects of combining two or more antimicrobics. In addition to increased
cost, there is increased risk of toxicity. Also, antagonism of antibacterial
eﬀectiveness may result when bacteriostatic and bactericidal agents are given
concurrently. Although antagonism of one antimicrobic by another has been
a frequent observation in vitro, documented clinical examples of this
phenomenon in veterinary medicine are unknown to us.

In a study evaluating antimicrobic susceptibility proﬁles in dogs over a 15-

year period (1984–1998), resistance to speciﬁc antimicrobic classes directly
correlated with apparent use of that antimicrobial drug. With the in-
troduction of ﬂuoroquinolones, resistant staphylococcal isolates increased
from 0% to 12%. During the same period, penicillin-type antibiotics were
prescribed less frequently and correlated with a decline in staphylococcal
isolates resistant to penicillin-type antibiotics. Results of two studies
evaluating bacteria isolated from skin of dogs also suggest that reductions
in resistance to particular antimicrobics paralleled decreased use of those
speciﬁc antibiotic classes

[13,14]

. These data and other information support

the premise that driving forces for emerging antimicrobial resistance are
repeated exposure of bacteria to antimicrobics (ie, selection pressure) and
access of bacteria to a large antimicrobic-resistant gene pool

. On the

basis of these observations, discontinuing antimicrobic therapy to determine
if persistent bacteria will become less resistant or be replaced by commensal
bacteria that are more susceptible to routine antimicrobials is sometimes
a rational therapeutic choice.

One question remains: ‘‘When is antimicrobic withdrawal safe for the

patent?’’ There are no clear guidelines on safety for discontinuing
antimicrobic therapy as a component of the therapeutic strategy for

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management of resistant bacterial UTIs in dogs and cats. These decisions
must be made on a case-by-case basis, balancing potential risks and
potential beneﬁts in conjunction with appropriate monitoring. When
considering withholding antimicrobic therapy, we attempt to predict the
patient’s ability to tolerate infection without further harm. Potential
sequelae to bacterial UTIs are listed in

Box 4

For example, although infections of the urinary bladder may cause

hematuria and dysuria, they are unlikely to cause systemic illness and are
typically associated with a low risk of mortality. Therefore, we are more
likely to consider discontinuing antimicrobic therapy for resistant infections
localized to the urinary bladder than resistant infections localized to the
kidneys. In a study of persistent and recurrent UTIs in dogs, more than half
were asymptomatic. In that same study, hematuria was absent in 53% and
pyuria was absent in 29%

. These ﬁndings indicate that when therapy of

some dogs with polyresistant bacterial UTI is withdrawn, these dogs are
unlikely to experience undue discomfort.

We discontinued therapy of several dogs and cats when uropathogens

developed resistance to antimicrobial therapy. Although these patients had
persistent signiﬁcant bacteriuria, they were asymptomatic (so-called ‘‘asymp-
tomatic bacteriuria’’). In some patients, the bacteria became less resistant to
antimicrobics after several months. In other patients, quantitative cultures
revealed that the original uropathogens were replaced by diﬀerent bacteria
that were less resistant. Some of these patients have remained asymptomatic
for several years without treatment. When one patient developed dysuria,
treatment with an antimicrobial drug was associated with rapid resolution of

Box 4. Potential sequelae to bacterial urinary tract infection

Lower urinary tract dysfunction

Dysuria, pollakiuria
Urge to incontinence
Damage to the detrusor muscle
Damage to the urethra

Prostatitis (acute or chronic)
Infertility
Struvite urolithiasis and its sequelae (especially urinary

obstruction)

Renal dysfunction (acute and chronic)

Pyelonephritis
Renal failure

Septicemia (especially in patients with concomitant obstruction

to urine flow)

Anemia of chronic infection

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clinical signs and elimination of the initial uropathogens (Proteus spp and
Escherichia coli

) from urine. Signiﬁcant numbers of Streptococcus spp were

cultured from urine collected on the seventh day of therapy (so-called
‘‘superinfection’’), however. These ﬁndings are not unique to dogs and cats.
In a prospective study of 50 older women randomized to receive therapy or no
therapy for asymptomatic UTI, there was no diﬀerence in morbidity or
mortality between the two groups over 12 months

. The group receiving

antibiotics had signiﬁcantly greater adverse drug reactions and development
of resistant reinfections, however.

If a patient is unresponsive to antimicrobic therapy and if diagnostic urine

cultures were not obtained before therapy, isolation of resistant bacteria in
subsequent cultures may indicate empiric selection of an ineﬀective antimi-
crobial drug. Some bacteria have intrinsic antimicrobic-resistant mechanisms
that are inherent to the species. For example, Pseudomonas spp are in-
trinsically resistant to most b-lactam antibiotics because they are suﬃciently
hydrophobic and these drugs do not diﬀuse through the outer bacterial
membrane. This situation could be avoided by initial selection of an
antimicrobic on the basis of in vitro susceptibility testing:

Culture Results

¼ New Bacterial Species

Cultures of urine during antimicrobial therapy that reveal eradication of

microbes identiﬁed by diagnostic cultures but growth of new uropathologic
bacteria indicate that previous antimicrobic therapy was eﬀective against the
initial pathogen. In this situation, it is apparent that the antimicrobial drug
predisposed the patient to infection with a resistant uropathogen (ie,
superinfection). Superinfections are deﬁned as infections with an additional
organism during the course of antimicrobial treatment. They are most likely
to occur in association with indwelling urethral catheters or as sequelae to
urinary diversion techniques in which the urinary tract communicates with
the intestinal tract. They also occur when proximal portions of the urethra,
urinary bladder, or kidneys communicate directly with the exterior (eg,
antepubic urethrostomy, tube cystostomy, percutaneous nephropyelos-
tomy). Unless the breach in host defenses can be identiﬁed and corrected,
recurrent infections are imminent even with eﬀective antimicrobic drugs to
treat the current infection. Because ongoing antimicrobic therapy is selecting
for resistant strains, it should be discontinued. To minimize drug-induced
superinfections, appropriate therapy should be directed toward eliminating
or controlling abnormalities in host defenses.

How to interpret urine cultures to diagnose and manage recurrent infections

Recurrence of clinical or laboratory signs of UTI after withdrawal of

therapy may be classiﬁed as bacterial relapse or reinfection. After successful

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antimicrobic therapy, timely follow-up evaluations (so-called ‘‘surveillance
cultures’’) are indicated to detect recurrent infections at a subclinical state
(see

http://www.aahanet.org/members_only/practice/02-Compliance.pdf

). Relapses (persistent infections) are deﬁned as recurrences caused

by the same species and serologic strain of bacteria, usually within several
weeks of cessation of therapy. Recovering the same organism from pre-
viously sterile urine is presumptive evidence that antimicrobial therapy failed
to eradicate the infection completely. Deep-seated infections or a lapse in
compliance is a common cause for bacterial relapses. For example, infections
of the canine prostate gland are a common cause for recurrent infections. In
this situation, relapses seem to be related to poor penetration of antimicro-
bial drugs into prostatic secretions. Reinfections may also be associated with
persistent abnormalities in prostatic defenses against bacterial infections.
Another cause of relapsing bacterial UTI may be linked to the recent
observation that uropathogenic E coli in mice can invade uroepithelial cells
and encase themselves in protective polysaccharide-rich pods that evade
immune stimulation and antimicrobic killing

[17,18]

. To eradicate relapsing

infections, consider treatment for a longer period (3–6 weeks) with
appropriate antimicrobics capable of greater tissue penetration (lipid-soluble
agents with low protein binding). To ensure that a suﬃcient concentration of
drug is maintained at the site of infection, administer medications at the
higher end of the dosage range (particularly when administering concentra-
tion-dependent antimicrobics, such as ﬂuoroquinolones) and with increased
frequency (particularly when administering time-dependent antimicrobics,
such as potentiated penicillins). Reculturing urine during therapy and after
therapy is essential to monitor therapeutic eﬃcacy.

Reinfections are deﬁned as recurrent infections caused by a diﬀerent

pathogen(s). If tissues of the urinary tract have had time to heal from
infection, reinfections often occur at a longer interval after cessation of
therapy than do relapses. Detection of frequent reinfections after antimi-
crobial therapy is an indication to evaluate the patient for predisposing
causes capable of breaching normal host defenses. Reinfections should be
managed by choosing antimicrobial agents on the basis of antimicrobial
susceptibility tests. Elimination of bacterial pathogens associated with
reinfections may require therapy of shorter duration (10–14 days) than
recurrences associated with relapses (3–6 weeks). Infrequent recurrences
(two or three times per year) may be treated as single episodes (ie, short
course of a suitable antimicrobial agent).

In some patients with recurrent reinfections, elimination of predisposing

causes may be impossible. In such cases, it may be helpful to provide low-
dose (preventative) antibacterial therapy for an indeﬁnite period (6 months
or more) with drugs primarily eliminated in urine. Reduced dosages
(approximately 33% to 50% of the daily therapeutic dose) of drugs excreted
in high concentration in urine may be used provided there has been
complete eradication of bacterial pathogens by therapeutic dosages of
appropriate drugs ﬁrst. Logically, preventative antimicrobial therapy would

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be inappropriate for management of patients with recurrent bacterial UTI
caused by relapses (reinfection with the same organism), because the
organism has never been eliminated.

Even though this preventative dosage regimen may not result in

bactericidal concentrations of antimicrobials throughout the day, low
concentrations of some drugs apparently interfere with development of
ﬁmbriae by some uropathogens. This, in turn, interferes with the ability of
potential pathogens to adhere to uroepithelial cells. It is best to give one
daily preventative dose of the antibiotic at a time when the drug is likely to
be retained in the urinary tract for several hours (ie, before bedtime).

During preventative therapy, urine samples collected by cystocentesis

(not by catheterization or voiding) should be recultured approximately once
each month. If bacteria are identiﬁed, a ‘‘breakthrough’’ infection may have
occurred. The patient should be treated again with therapeutic dosages of an
antimicrobial drug selected on the basis of susceptibility tests. Once the
infection has been eradicated and the associated inﬂammatory response
subsides (usually 2–4 weeks), preventative therapy may be resumed.

After 6 to 9 months of consecutive negative urine cultures, therapy may

be discontinued on a trial basis to determine if a reinfection will occur. If
abnormalities in host defenses have healed, UTI may not recur. If UTI
develops within a short period, the procedures outlined previously should be
repeated.

Long-term use of antimicrobial agents is not without risk of adverse

eﬀects. For example, sulfadiazine-trimethoprim combinations have been
associated with keratoconjunctivitis sicca, folate deﬁciency anemia, and
immune complex reactions. The potential for long-term low-dose antimi-
crobic therapy to contribute to bacterial resistance in commensal bacteria in
dogs and cats is unknown but appears likely.

Summary

Quantitative urine cultures and antimicrobial susceptibility proﬁles pro-

vide an accurate method of diagnosing bacterial infection and monitoring
therapy. Diagnostic urine cultures are performed prior to initiation of
antimicrobic therapy and are essential to avoid treating noninfectious
disorders with antimicrobial drugs. Therapeutic urine cultures are obtained
during antimicrobic therapy and are used to verify antimicrobic eﬃcacy and
to help determine reasons for antimicrobic failure. Surveillance urine
cultures are preformed shortly after completion of successful antimicrobic
therapy. Surveillance cultures are used to detect recurrent infections at
a subclinical state and are essential to diﬀerentiate relapses from reinfections.

References

[1] Ling GV. Therapeutic strategies involving antimicrobic treatment of the canine urinary

tract. J Am Vet Med Assoc 1984;185:1162–4.

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J.P. Lulich, C.A. Osborne / Vet Clin Small Anim 34 (2004) 1027–1041

[2] Kivisto AK, Vasenius H, Sandholm M. Canine bacteriuria. J Small Anim Pract 1977;18:

707–12.

[3] Norris CR, Williams BJ, Ling GV, et al. Recurrent and persistent urinary tract infections

in dogs: 383 cases (1969–1995). J Am Anim Hosp Assoc 2000;36:484–92.

[4] Seguin MA, Vaden SL, Altier C, et al. Persistent urinary tract infections and reinfections in

100 dogs (1989–1999). J Vet Intern Med 2003;17:622–31.

[5] Goett SD, Degner DA. Persistent urinary problems in a young golden retriever. Vet Med

2003;98:839–42.

[6] American Animal Hospital Association. Understanding the compliance gap. Available at:

. Accessed December

3, 2003.

[7] Ling GV. Treatment of urinary tract infections with antimicrobial agents. In: Kirk RW,

editor. Current veterinary therapy VIII, small animal practice. Philadelphia: WB Saunders;
1983. p. 1051–5.

[8] Scheiner J, Altemeier WA. Experimental study of factors inhibiting absorption and

eﬀective therapeutic levels of declomycin. Surg Gynecol Obstet 1962;114:9–14.

[9] Oluoch A, Weisiger R, Siegel AM, Campbell KL, Krawiec DR, McKiernan BC. Trends of

bacterial infections in dogs: characterization of Staphylococcus intermedius isolates (1990–
1992). Canine Pract 1996;21:12–9.

[10] Fish DN, Choi MK, Jung R. Synergic activity of cephalosporins plus ﬂuoroquinolones

against Pseudomonas aeruginosa with resistance to one or both drugs. J Antimicrob
Chemother 2002;50:1045–9.

[11] Karlowsky JA, Zelenitsky SA, Zhanel GG. Aminoglycoside adaptive resistance.

Pharmacotherapy 1997;17:549–55.

[12] Klastersky J. Empirical treatment of sepsis in neutropenic patients. Int J Antimicrob

Agents 2000;16:131–3.

[13] Kruse H, Hofshagen M, Thoresen SI, et al. The antimicrobial susceptibility of

staphylococcus species isolated from canine dermatitis. Vet Res Commun 1996;20:205–14.

[14] Lloyd DH, Lamprot AI, Feeney C. Sensitivity to antibiotics amongst cutaneous and

mucosal isolates of canine pathogenic staphylococci in the UK, 1980–96. Vet Dermatol
1996;7:171–5.

[15] Hoﬀman SB. Mechanisms of antibiotic resistance. Compend Contin Educ Pract Vet 2001;

23:464–73.

[16] Nicolle LE, Mayhew WJ, Bryan L. Prospective randomized comparison of therapy and no

therapy for asymptomatic bacteriuria in institutionalized elderly women. Am J Med 1987;
83:27–33.

[17] Anderson GG, Palermo JJ, Schilling JD, et al. Intracellular bacterial bioﬁlm-like pods in

urinary tract infections. Science 2003;301:105–7.

[18] Mulvey MA, Lopez-Boada YS, Wilcon CL, et al. Induction and evasion of host defenses

by type 1-piliated uropathogenic Escherichia coli. Science 1998;282:1494–7.

1041

J.P. Lulich, C.A. Osborne / Vet Clin Small Anim 34 (2004) 1027–1041

Feline idiopathic cystitis: current

understanding of pathophysiology

and management

Jodi L. Westropp, DVM,

C.A. Tony Buﬃngton, DVM, PhD*

The Ohio State University Veterinary Hospital, 601 Tharp Street, Columbus,

OH 43210–1089, USA

Signs referable to the lower urinary tract of indoor-housed cats have been

described in the veterinary literature for at least 80 years

. The terms feline

urologic syndrome

(FUS) and feline lower urinary tract disease (FLUTD)

were coined in the 1970s (FUS

) and 1980s (FLUTD

) to describe

variable combinations of straining, hematuria, pollakiuria (frequent passage
of small amounts of urine), and periuria (urinations in inappropriate
locations) seen in cats with the disorder. No diagnosis for clinical signs of
irritative voiding can be determined in approximately two thirds of cats with
lower urinary tract (LUT) signs, so we refer to them as having feline
idiopathic cystitis (FIC). If cystoscopy is performed and characteristic
submucosal petechial hemorrhages are seen, a diagnosis of feline interstitial
cystitis can be made

. This term was chosen because of similarities

between cats and human beings with interstitial cystitis, an idiopathic pelvic
pain syndrome that is characterized by diﬃcult, painful, and frequent
urinations without a diagnosable cause

Pathophysiology

Two forms of interstitial cystitis are recognized in human beings, the

common nonulcerative and uncommon ulcerative forms

. Most cats with

feline interstitial cystitis have signs comparable to the nonulcerative form,
although the ‘‘Hunner’s ulcers’’ sometimes seen in human beings with
interstitial cystitis have been described in a cat

. The term feline idiopathic

* Corresponding author.
E-mail address:

buﬃngton.1@osu.edu

(C.A. Tony Buﬃngton).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.002

Vet Clin Small Anim

34 (2004) 1043–1055

cystitis

is used in this article, because cats with this disorder do not

commonly undergo cystoscopic evaluation (and most do not need to be
subjected to this procedure).

Based on recent research we believe that FIC may include multiple

complex abnormalities of the nervous and endocrine systems that likely
aﬀect more than just the urinary bladder

. Enhanced central noradren-

ergic drive in the face of inadequate adrenocortical restraint seems to be
related to maintaining the chronic disease process (

). These systems

seem to be driven by tonically increased hypothalamic corticotropin-
releasing factor release, which may represent the outcome of a developmen-
tal accident

[8,9]

. Because of these abnormalities, treatment strategies that

decrease central noradrenergic drive may be important in reducing signs of
FIC; those that do not address this aspect of the disease seem to be less
eﬀective. Until more eﬀective treatments to normalize responsiveness of the
stress response system are available, eﬀorts to reduce input to this system by
environmental enrichment seem reasonable

[10,11]

Brainstem

nuclei

Neurosteroids

Hypothalamus

Pituitary

SNS

Bladder,

other organs

ACTH

Adrenal

Cortex

CRF

Cortisol

CNS

Inhibition

Excitation

Fig. 1. Imbalanced neuroendocrine system of cats with feline idiopathic cystitis. Excitatory
sympathetic nervous system (SNS) outﬂow is inadequately restrained by cortisol. This enhanced
activity can increase tissue permeability, resulting in increased sensory aﬀerent activity.
Feedback inhibition at the level of the anterior pituitary and hypothalamus also is reduced,
which tends to perpetuate corticotrophin-releasing factor (CRF) output. Neurosteroid
production by the adrenal cortex, which generally enhances central nervous system (CNS)
inhibitory tone during chronic stress, also may be reduced. The solid lines indicate stimulation,
and the dotted lines indicate inhibition. Line thickness is intended to indicate intensity of the
signal.

1044 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

Environmental enrichment

Just as water is primary therapy for prevention of urinary stone

recurrence, we think that environmental enrichment is primary therapy
for prevention of recurrence of elimination disorders, including FIC. This
opinion is based on documented neuroendocrine abnormalities suﬀered by
cats with FIC

[8,12,13]

and on our clinical experience. We deﬁne environ-

mental enrichment for indoor-housed cats to mean provision of all
‘‘necessary’’ resources, reﬁnement of interactions with owners, a tolerable
intensity of conﬂict, and thoughtful institution of change(s). Although we
are not aware that a particular resource list has been validated for indoor-
housed cats, some recommendations are available

[11,14,15]

. We have

assembled a provisional list in

that could be used by owners to guide

consideration of these parameters for each of their cats. The following
tentative recommendations are organized to follow

; more compre-

hensive suggestions are available in the many excellent publications

about cat housing and behavior that currently are available. We also
recommend extending the ‘‘1

þ 1’’ rule traditionally applied to litter boxes

(one for each cat in the home plus one more) to all pertinent resources
(particularly food, water, and litter containers) in the household.

Food

Behavioral and ethologic research suggest that cats prefer to eat

individually in a quiet location, where they are not startled by other
animals, sudden movement, or activity of an air duct or appliance that may
begin operation unexpectedly

[17,18]

. Although canned food may be

preferable for cats with FIC because of increased water content or a more
natural ‘‘mouth feel,’’ some cats may prefer dry foods. The phytoestrogen
content of soy used in some cat diets also could conceivably play a role in
modulating discomfort associated with the disease

. If a diet change is

appropriate, oﬀering the new diet in a separate adjacent container rather
than removing the usual food and replacing it with new food permits the cat
to express its preference. Natural cat feeding behavior also includes
predatory activities, such as stalking and pouncing. These may be simulated
by hiding small amounts of food around the house or by putting dry food in
a container from which the cat has to extract individual pieces or move
something to release the food pieces if such interventions appeal to the cat.
Also, some cats seem to have speciﬁc prey preferences. For example, some
cats prefer to catch birds, whereas others may prefer to chase mice or bugs.
Identifying a cat’s ‘‘prey preference’’ allows one to buy or make toys that the
cat is more likely to play with. Prey preference can be identiﬁed by paying
close attention to the cat’s reaction to toys with speciﬁc qualities, such as
those that resemble birds (feather toys), small mammals (‘‘furry mice’’), or
insects (laser pointers, pieces of dry food) presented one at a time or
together.

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Table 1
Environmental survey for indoor-housed cats

Yes

Food and water

Each cat has its own food and water bowl in a convenient location that provides

some privacy while eating or drinking and an ‘‘escape’’ route

Bowls are located such that another animal cannot sneak up on the cat while it

eats

Bowls are located away from appliances or air ducts that could come on

unexpectedly while the cat eats or drinks

Food and water is kept fresh (daily)
Bowls are washed regularly (at least weekly) with a mild detergent
The brand or type of food purchased is changed infrequently (less than monthly)
If a new food is oﬀered, it is put in a separate dish next to the familiar food so the

cat can choose to eat it if it wants to

Litter box management

Boxes are located on more than one level in multilevel houses
Boxes are located so another animal cannot sneak up on the cat while it uses one
Boxes are located away from appliances or air ducts that could come on

unexpectedly while the cat uses one, and an ‘‘escape’’ route is provided

The litter is kept clean and scooped as soon after use as possible (just like we

ﬂush after each use), at least daily

Boxes are washed regularly (at least weekly) with a mild detergent (like

dishwashing liquid) rather than strongly scented cleaners

Unscented clumping litter is used
The brand or type of litter purchased is changed infrequently (less than monthly)
If a new type of litter is oﬀered, it is put in a separate box so the cat can choose to

use it if it wants to

Each cat has its own litter box in a convenient well-ventilated location that still

gives the cat some privacy while using it

Environmental considerations

Scratching posts are provided
Toys are provided, rotated, or replaced regularly
Each cat has the opportunity to move to a warmer or cooler area if it chooses to
Each cat has a hiding area where it can get away from threats if it chooses to
Each cat has its own space that it can use if it chooses to

Rest

Each cat has its own resting area in a convenient location that still provides some

privacy, and an ‘‘escape’’ route

Resting areas are located such that another animal cannot sneak up on the cat

while it rests

Resting areas are located away from appliances or air ducts that could come on

unexpectedly while the cat rests

If a new bed is provided, it is placed next to the familiar bed so the cat can choose

to use it if it wants to

Movement

Each cat has the opportunity to move about freely, explore, climb, stretch, and

play if it chooses to

Social contact

Each cat has the opportunity to engage in play with other animals or the owner if

it chooses to

1046 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

Water

Cats also seem to have preferences for water that can be investigated.

Water-related factors to consider include freshness, taste, movement (water
fountains, dripping faucets, or aquarium pump-bubbled air into a bowl),
and shape of the container (some cats seem to resent having their vibrissae
touch the sides of the container when drinking). As with foods, changes in
water-related factors should be oﬀered in such a way that permits the cat to
express its preferences. Additionally, food and water bowls should be
cleaned regularly unless individual preference suggests otherwise.

Litter boxes

Litter boxes should be provided in diﬀerent locations throughout the

house to the extent possible, particularly in multiple cat households

Placing litter boxes in quiet convenient locations that provide an escape
route if necessary for the cat could help to improve conditions for normal
elimination behaviors. If diﬀerent litters are oﬀered, it may be preferable to
test the cat’s preferences by providing them in separate boxes, because
individual preferences for litter type have been documented. For cats with
a history of urinary problems, unscented clumping litter should be
considered. Litter boxes should be cleaned regularly and replaced; some
cats seem quite sensitive to dirty litter boxes. Litter box size and whether or
not it is open or covered also may be important to some cats.

Space

Cats interact with physical structures and other animals, including

human beings, in their environment. The physical environment should
include opportunities for scratching (horizontal and vertical options may be
necessary), climbing, hiding, and resting. Cats seem to prefer to monitor
their surroundings from elevated vantage points, so climbing frames,
hammocks, platforms, raised walkways, shelves, or window seats may
appeal to them. Playing a radio to habituate cats to sudden changes in
sound and human voices also has been recommended, and videotapes to
provide visual stimulation are available.

Play

Some cats seem to prefer to be petted and groomed, whereas others may

prefer play interactions with owners. Cats also can be easily trained to
perform behaviors (‘‘tricks’’); owners just need to understand that cats
respond much better to praise than to force and seem to be more amenable
to learning if the behavior is shaped before feeding. Cats also may enjoy
playing with toys, particularly those that are small, move, and mimic prey
characteristics. Many cats also prefer novelty, so a variety of toys should be
provided and rotated or replaced regularly to sustain their interest.

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Conﬂict

When cats’ perception of safety becomes threatened, they appear to

respond in an attempt to restore their ‘‘perception of control.’’ During
such responses, some cats become aggressive, some become withdrawn,
and some become ill. In our experience, intercat conﬂict commonly is
present when multiple cats are housed indoors together and health
problems exist. Conﬂict among cats can develop because of threats to
their perception of their overall status or rank in the home, from other
animals in the home, or from outside cats

[21]

. With a little practice, one

can recognize signs of conﬂict and estimate its potential role in
exacerbating signs of FIC. If it is, owners usually can identify causes
after signs of conﬂict are explained to them. Once this has been done,
clients often are well on their way to reducing intensity of conﬂict. Of
course, some conﬂict between housemates is normal, regardless of species.
Our goal is to reduce unhealthy conﬂict to a more manageable level for
cats involved.

Signs of conﬂict (

Table 2

) between cats can be open or silent. Signs of

open conﬂict are easy to recognize; the cats may stalk each other, hiss, and
turn sideways with legs straight and hair standing on end to make
themselves look larger. If neither cat backs down, the displays may increase
to swatting, wrestling, and biting. The signs of silent conﬂict can be so subtle
that they are easily missed. The cat creating conﬂict (assertive cat) can be
identiﬁed as the one that never backs away from other (threatened) cats,
denies other cats’ access to resources, stares at other cats, and lowers its
head and neck while elevating its hindquarters as it approaches less
conﬁdent cats. Hair along its back and on its tail and tail base may stand
on end, although not to the extent of cats engaged in open conﬂict, and the
cat may emit a low growl. The assertive cat eventually may only have to
approach or stare at a threatened cat for it to leave a resource, such as food
or a litter box. If the threatened cat tries to use the resource later, the
assertive cat’s presence alone may be enough to make it ﬂee. Because cats do
not seem to possess distinct dominance hierarchies or conﬂict resolution
strategies, threatened cats may attempt to circumvent agonistic encounters
by avoiding other cats, by decreasing their activity, or both. Threatened cats
often spend increasingly large amounts of time away from the family,
staying in areas of the house that others do not use, or they attempt to
interact with family members only when the assertive cat is elsewhere.

Signs of conﬂict can result from two types of conﬂict: oﬀensive and

defensive. In oﬀensive conﬂict situations, the assertive cat moves closer to
other cats and controls the interaction. In defensive conﬂict situations,
a threatened cat attempts to increase distance between itself and the thing it
perceives as a threat. Although cats engaged in either type of conﬂict may
spray or eliminate outside the litter box, we ﬁnd that threatened cats are
more likely to develop elimination problems.

1048 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

The most common cause of conﬂict between indoor-housed cats that we

have been able to identify is competition for resources. Cats may engage in
open or silent conﬂict over space, food, water, litter boxes, perches, sunny
areas, safe places where the cat can watch its environment, or attention
from people. There may be no obvious limitation to access to these
resources for conﬂict to develop. The change may only be the cat’s
perceptions of how much control it wants over the environment or its
housemates’ behaviors.

Open conﬂict is most likely to occur when a new cat is introduced into

the house and when cats that have known each other since kittenhood
reach social maturity. Conﬂict occurring when a new cat is introduced is
easy to understand, and good directions are available from many sources
for introducing the new cat to the current residents. Clients may be
puzzled by conﬂict that starts when one of their cats becomes socially
mature or when a socially mature cat perceives that one of its house-
mates is becoming socially mature. Cats become socially mature between
2 and 5 years of age and start to take some control of social groups and
their activities. This may lead to open conﬂict between male cats, between
female cats, or between male and female cats. Although clients may be
surprised, ‘‘because they lived so well together for the ﬁrst few years of
their lives,’’ a cat’s perceptions of resource needs may expand with social
maturity.

Cats that are familiar with each other but unevenly matched often show

conﬂict in more subtle ways. One of the cats in the conﬂict asserts itself,
and another cat is threatened by this cat’s actions. Silent conﬂicts may
not even be recognized until the threatened cat begins to hide from the

Table 2
Signs of silent conﬂict between cats

Assertive cat

Threatened cat

Never backs away from other cats

Spends large amounts of time

hiding or away from the family

Stares at other cats

Avoids eye contact with other cats

Denies other cats access resources

Yields resources to other cats

Rub cheeks, head, chin, and

tail on people, doorways, and
furniture at cat height

When it sees the threatened cat

When it sees the assertive cat

Lowers its head and neck while

elevating its hindquarters and stalks
the other cat

Crouches, may cower, may then ﬂee

Piloerects the hair along its back, tail

base, and tail

Growls

Does not vocalize

May spray

May spray
May develop cystitis or other

disease problem

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J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

assertive cat, starts to hiss or ﬁght back when it sees the other cat, or
develops a health problem.

In addition to signs of conﬂict described previously, the assertive cat

can be identiﬁed by its marking behavior. These cats rub their cheeks,
head, chin, and tail on people, doorways, and furniture at cat height.
Unfortunately, silent conﬂict can also involve urine, including marking by
the assertive or threatened cat and cystitis in the threatened cat. Conﬂict-
related urine marking can include spraying, where the cat treads and
kneads, raises its tail, and ﬂicks the tip of it while spraying urine on
a vertical surface or squatting and urinating outside the litter box
(nonspray marking). Both male and female cats may spray, and although
neutering reduces frequency of spraying, it cannot eliminate the behavior.
Conﬂict-related urine marking can be exhibited by either the assertive or
threatened cat, but in our experience, FIC usually occurs in the
threatened cat; we have even seen threatened male cats spray bloody
urine. Cats that urinate on bedspreads or other elevated open places may
do so because their access to the litter box is restricted by another cat or
if they are afraid to use the box because it is placed such that a quick
escape from another cat might not be possible.

Treatment for conﬂict between cats involves providing a separate set of

resources for each cat, preferably in locations where the cats can use them
without being seen by other cats. This lets cats avoid each other if they
choose to without being deprived of an essential resource. Conﬂict also can
be reduced by neutering all the cats and by keeping all nails trimmed as
short as practicable. Whenever cats involved in the conﬂict cannot be
directly supervised, they may need to be separated. This may mean that
some of the cats in the household can stay together but that the threatened
cat is provided a refuge from the other cats. This room should contain all
necessary resources for the cat staying in it.

Cats generally require and use more space than the average house or

apartment aﬀords them. Addition of elevated spaces such as shelves, ‘‘kitty
condos,’’ cardboard boxes, beds, or crates may provide enough space to
reduce conﬂict to a tolerable level. In severe situations, some cats may
beneﬁt from behavior-modifying medications. In our experience, however,
medication can help only after environmental enrichment has occurred; it
cannot replace it.

Cats involved in the conﬂict may never be ‘‘best friends,’’ but they usually

can live together without showing signs of conﬂict or conﬂict-related disease.
In severe cases, a behaviorist may be consulted for assistance in desensitizing
and counterconditioning of cats in conﬂict so they can share the same spaces
more comfortably if this is desired.

Conﬂict with other animals, dogs, children, or adults is relatively

straightforward. In addition to being solitary hunters of small prey, cats
are small prey themselves for other carnivores, including dogs. Regardless of
how sure the client is that his or her dog will not hurt the cat, to the cat, the

1050 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

dog represents a predator. If the cat does not assert dominance over the dog,
as often happens, it must be provided ways to escape at any time. For
human beings, it usually suﬃces to explain that cats may not understand
rough treatment as play but as a predatory threat.

Most cats in urban areas in the United States are housed indoors and

neutered, so conﬂict with outside cats can occur when a new cat enters the
area around the house the aﬀected cat lives in. To cats, windows are no
protection from a threatening cat outside. If outside cats are the source
of the problem, a variety of strategies to make one’s garden less desirable
to them are available.

Because of the dearth of controlled trials, it currently is not possible to

prioritize the importance of any of these suggestions or to predict which
would be most appropriate in any particular situation. Appropriately
designed epidemiologic studies might be able to identify particularly
important factors, after which intervention trials could be conducted to
determine their eﬃcacy in circumstances in which owners successfully
implemented suggested changes. Our recent clinical experience includes 76
indoor-housed cats with recurrent FIC we have been following to attempt
to determine eﬀects of environmental enrichment during a 1-year period;
17 are individually housed, and 59 live in multicat households. There are
49 male cats and 27 female cats, all neutered, ranging in age from 1 to 10
years. Based on structured interviews at the time of referral, the most
common cause of clinical signs in the singly housed cats was separation
anxiety; for the cats from multiple cat households, the most common
cause of clinical signs was some form of conﬂict. At the time of this
writing, 19 cats have completed the study; median time of follow-up for
the 57 cats currently being studied is 6 months. In the singly housed cats,
a single episode of recurrence has been observed in 2 cats (12%
recurrence rate). In the cats in multiple cat households, recurrences have
been reported in 7 cats (12% recurrence rate). Published recurrence rates
in the absence of intervention are in the range of 50%. In all these cats,
the primary chronic therapy intentionally instituted was environmental
enrichment to reduce central neural arousal. These results suggest that
expression of FIC may result from the presence of a susceptible cat in
a provocative environment. Our challenge is to identify and change what
the susceptible cat is threatened by to prevent future bouts. Moreover,
only about half of clients oﬀered this approach are amenable to it, and
we have not conducted a formal intention-to-treat analysis of the results
because the study is still in progress. Nevertheless, results are encourag-
ing, and we have not encountered adverse eﬀects related to this approach.

Additional approaches

Once environmental enrichment strategies have been implemented,

additional treatments may be considered. In our experience, these

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approaches are more likely to succeed after the environment has been en-
riched to the extent possible by the client and more likely to fail in the
absence of environmental enrichment. They are listed in the order in which
we consider them.

Pheromones

A novel aspect of environmental enrichment that recently has become

available is the application of pheromones to the living space

Pheromones are chemical substances that seem to transmit highly speciﬁc
information between animals of the same species. Although the exact
mechanism of action is unknown at this time, pheromones seem to eﬀect
changes in the function of the limbic system and the hypothalamus to alter
the emotional status of animals. Five facial pheromones have been isolated
from cats; cats deposit the F3 fraction on prominent objects (including
human beings) by rubbing against the object when the cat feels safe and at
ease. The function of this secretion is not only to mark objects but to serve
as an antagonist for urine marking and scratching.

Feliway (Farnum Companies Inc., Phoenix, AZ) a synthetic analogue of

this naturally occurring feline facial pheromone, was developed to decrease
anxiety-related behaviors of cats. Although not speciﬁcally tested in cats with
FIC, treatment with this pheromone has been reported to reduce the amount
of anxiety experienced by cats in unfamiliar circumstances, a response that
may be helpful to these patients and their owners. Decreased spraying in
multicat households, decreased marking, and a signiﬁcant decrease in
scratching behavior also have been reported subsequent to its use. Although,
Feliway is not a panacea for unwanted cat behaviors or FIC, we have used it
successfully in combination with environmental enrichment or drug therapies.

Feliway is sold as a spray and as a room diﬀuser. The spray can be used

to treat areas of the house where the cat is urinating by use of a single spray
to the aﬀected spot for 30 days. We also have found Feliway to be beneﬁcial
to decrease anxiety associated with traveling. Clients can spray the cat
carrier at least 15 minutes before the trip and then place the cat in the carrier
to help decrease the stress and anxiety that most cats associate with travel.
Treated areas should be sprayed at least 15 minutes before the cat en-
counters the area, because the vehicle (ethanol) in which the pheromone is
carried is oﬀensive to most cats. The room diﬀuser can be placed in a room
where the cat inappropriately urinates. One room diﬀuser is reported to
cover approximately 650 square feet and to last for 30 days. This method of
administering pheromone is new, and we have little experience with its use.

Analgesia

We provide short-term analgesic therapy in cases of an acute ﬂare of signs

as appropriate and sometimes recommend pharmacotherapy in refractory
cases after all previous recommendations have been instituted to the extent

1052 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

possible, because we have not found a drug that is more eﬀective than
environmental enrichment. Drugs that might be considered include tricyclic
‘‘antidepressants’’ and other autonomic nervous system modulators, gly-
cosaminoglycan-replacing compounds, sedatives, gamma-aminobutyric ac-
id-A (GABA

) agonists, and neuroactive steroids. Many aspects of this

therapy, which is rapidly evolving, have not been adequately tested to
permit evidence-based recommendations. Because placebo response also
may be mediated by a reduction in this drive, diﬀerentiating placebo eﬀects
from drugs eﬀects can be diﬃcult. In most studies of human patients with
interstitial cystitis, the placebo response rate is in the range of 50%.

Treating the owner

Finally, decreasing central neural drive in the cat may require reducing the

level of arousal of the owner to the situation, as has been reported in human
beings (parents and children

). We try to follow recommendations

http://www.nssvet.org/ici/

for

improving client satisfaction with our suggestions by attempting to ensure
that clients leave the encounter with us feeling that they:

1. Were listened to
2. Received an explanation for the problem that made sense to them
3. Felt care and concern being expressed by the caregivers and others in the

clinic

4. Left with an enhanced sense of mastery or control over the cat’s illness

or its signs

Because FIC can be a chronic frustrating disease, we have found that

keeping these four points in mind when communicating with clients is
beneﬁcial for the client, pet, and clinician. We have also established
a technician program in which a staﬀ member follows only these patients
as often as necessary to be sure their cat’s problems are explained
thoroughly and the clients gain enough understanding of the disease process
to feel comfortable with managing their cat’s disease.

Once clients have identiﬁed areas for improvement in resource availabil-

ity, they may need help in instituting changes. In our experience, veterinary
technicians are invaluable in helping clients with all aspects of this change
process, because they often have more time available to answer questions
and provide guidance. We also recommend that environmental modiﬁca-
tions be instituted slowly, one at a time, and in a way that permits the cat to
express its like or dislike for the change. For example, new diets or litter
should be oﬀered in separate containers next to the usual food or litter so
the cat can express a preference.

Summary

Many indoor-housed cats seem to survive perfectly well by accommo-

dating to less than perfect surroundings. Neuroendocrine abnormalities in

1053

J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

the cats we treat, however, do not seem to permit adaptive capacity of
healthy cats, so these cats may be considered a separate population with
greater needs. Moreover, veterinarians are concerned more with optimizing
environments of indoor cats than with identifying minimal requirements for
indoor survival. Further information about environmental enrichment and
conﬂict is available at:

References

[1] Kirk H. Retention of urine and urine deposits. In: The diseases of the cat and its general

management. London: Bailliere, Tindall and Cox; 1925. p. 261–7.

[2] Osbaldiston GW, Taussig RA. Clinical report on 46 cases of feline urological syndrome.

Vet Med Small Anim Clin 1970;65:461–8.

[3] Osborne CA, Johnston GR, Polzin DJ, et al. Redeﬁnition of the feline urologic syndrome:

feline lower urinary tract disease with heterogeneous causes. Vet Clin North Am Small
Anim Pract 1984;14:409–38.

[4] Buﬃngton CAT, Chew DJ, Woodworth BE. Feline interstitial cystitis. J Am Vet Med Assoc

1999;215:682–7.

[5] Buﬃngton CAT, Chew DJ, Woodworth BE, et al. Idiopathic cystitis in cats: an animal

model of interstitial cystitis. In: Sant GR, editor. Interstitial cystitis. Philadelphia:
Lippincott-Raven; 1997. p. 25–31.

[6] Sant GR, editor. Interstitial cystitis. Philadelphia: Lippincott-Raven; 1997. p. 284.
[7] Clasper M. A case of interstitial cystitis and Hunner’s ulcer in a domestic shorthaired cat.

NZ Vet J 1990;38:158–60.

[8] Westropp JL, Welk KA, Buﬃngton CAT. Small adrenal glands in cats with feline

interstitial cystitis. J Urol 2003;170:2494–7.

[9] Osadchuk LV, Braastad BO, Hovland AL, et al. Handling during pregnancy in the blue

fox (Alopex lagopus): the inﬂuence on the fetal pituitary-adrenal axis. Gen Comp
Endocrinol 2001;123:100–10.

[10] Morley-Fletcher S, Rea M, Maccari S, et al. Environmental enrichment during adolescence

reverses the eﬀects of prenatal stress on play behaviour and HPA axis reactivity in rats. Eur
J Neurosci 2003;18:3367–74.

[11] Laule GE. Positive reinforcement training and environmental enrichment: enhancing

animal well-being. J Am Vet Med Assoc 2003;223:969–73.

[12] Reche AJ, Buﬃngton CAT. Increased tyrosine hydroxylase immunoreactivity in the locus

coeruleus of cats with interstitial cystitis. J Urol 1998;159:1045–8.

[13] Buﬃngton CAT. Plasma catecholamine concentrations in cats with interstitial cystitis

[abstract]. J Urol 2000;163:58.

[14] Buﬃngton CAT. External and internal inﬂuences on disease risk in cats. J Am Vet Med

Assoc 2002;220:994–1002.

[15] Bracke MBM, Spruijt BM, Metz JHM. Overall animal welfare reviewed. Part 3: welfare

assessment based on needs and supported by expert opinion. Neth J Agric Sci 1999;47:
307–22.

[16] Overall KL. Feline elimination disorders. In: Clinical behavioral medicine for small

animals. St. Louis (MO): Mosby; 1997. p. 160–94.

[17] Turner DC, Bateson P, editors. The domestic cat: the biology of its behaviour. 2nd edition.

Cambridge: Cambridge University Press; 2000. p. 244.

[18] Masserman JH. Experimental neuroses. Sci Am 1950;182:38–43.
[19] Shir Y, Campbell JN, Raja SN, et al. The correlation between dietary soy phytoestrogens

and neuropathic pain behavior in rats after partial denervation. Anesth Analg 2002;94:
421–6.

1054 J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

[20] Horwitz DF. Behavioral and environmental factors associated with elimination behavior

problems in cats: a retrospective study. Appl Anim Behav Sci 1997;52:129–37.

[21] Barry KJ, Crowell-Davis SL. Gender diﬀerences in the social behavior of the neutered

indoor-only domestic cat. Appl Anim Behav Sci 1999;64:193.

[22] Pageat P, Gaultier E. Current research in canine and feline pheromones. Vet Clin North

Am Small Anim Pract 2003;33:187.

[23] Webster-Stratton C, Herbert M. What really happens in parent training. Behav Modif

1993;17:407–56.

[24] Brody H. The placebo response—recent research and implications for family medicine.

J Fam Pract 2000;49:649–54.

1055

J.L. Westropp, C.A. Tony Buﬃngton / Vet Clin Small Anim 34 (2004) 1043–1055

Surgical management of urinary

incontinence

Michael G. Hoelzler, DVM,

David A. Lidbetter, BVSc, MVS*

Department of Small Animal Clinical Sciences, C247 Veterinary Teaching Hospital,

The University of Tennessee College of Veterinary Medicine Knoxville,

TN 37996–4544, USA

Surgery for urinary incontinence

Urinary incontinence occurs in dogs and cats. Female animals are more

frequently aﬀected. When presenting with incontinence, young animals are
more commonly aﬀected with ureteral ectopia, whereas older animals more
often suﬀer from urethral sphincter mechanism incompetence (USMI).
Surgery may be a useful tool to help improve or resolve both conditions.

Urethral sphincter mechanism incompetence

Pathophysiology

USMI is the most common cause of urinary incontinence in female dogs.

It is estimated that more than 20% of the general population of spayed
bitches are aﬀected as well as 30% of spayed bitches heavier than 20 kg

Approximately 75% of bitches become incontinent within 12 months of
ovariohysterectomy, and 28% become incontinent immediately after the
procedure

. Although spayed bitches with acquired disease predominate,

the disease can occur congenitally and is called juvenile USMI. Holt and
Thrusﬁeld

estimate that approximately 50% of juvenile incontinent

bitches recover continence after their ﬁrst estrus. Large and giant breeds
have increased risk, and small breeds have a low risk for development of the
disorder. Breeds at highest risk include the Old English Sheep Dog, Irish
Setter, Doberman Pinscher, and Rottweiler

* Corresponding author.
E-mail address:

lid@utk.edu

(D.A. Lidbetter).

0195-5616/04/$ - see front matter

doi:10.1016/j.cvsm.2004.03.003

Vet Clin Small Anim

34 (2004) 1057–1073

The exact etiology and pathogenesis of the disease have not been

elucidated. Multifactorial contributory factors include decreased urethral
pressure, reduced urethral length, possible estrogen deﬁciency, intrapelvic
bladder, deﬁcient vesicourethral support, and urethral smooth muscle
abnormalities

[3,5–7]

. Tail docking and obesity are also considered risk

factors

Clinical features

Bitches with USMI are able to void appropriately and consciously.

Incontinence is most often associated with relaxation or recumbency,
particularly at night. A typical historical ﬁnding is the owner discovering
a puddle of urine after the animal has been sleeping. Incontinence
deteriorates associated with urinary tract infection

Diagnosis

Pertinent historical ﬁndings, such as age at onset of incontinence and

frequency and pattern of inappropriate urination are critical to the working
diagnosis. Incontinence must be distinguished from polyuria and pollakiu-
ria. A thorough physical examination is performed as part of a minimum
database. The vulva and perineum are examined for signs of immaturity and
urine scalding or dermatitis, which can be associated with incontinence. The
bladder is palpated to try to assess wall thickness and the presence of masses
or calculi. Urination is observed, and an assessment of residual volume is
made.

Diagnosis can be conﬁdently made based on signalment, history, and

contrast

radiographs.

Quantitative

evaluation

requires

the

use

of proﬁlometry.

Urinalysis

Severe cystitis can result in involuntary bladder contractions and increased

urine leakage in some bitches

. When urinalysis is suggestive of cystitis,

culture and sensitivity testing should follow and the animal should be treated
with appropriate antibiotics for 3 weeks to see if the incontinence resolves.

Radiographs

Plain lateral and ventrodorsal abdominal radiographs are taken to help

conﬁrm the diagnosis of USMI. These also help to rule out other medical
conditions. A positive-contrast vaginourethrocystogram can be used to
examine the location of the bladder neck and to help deﬁne the bladder
shape. USMI bitches characteristically have an intrapelvic bladder neck (the
bladder neck is located caudal to the cranial rim of the pubis on a lateral
ﬁlm), which may have a brick-shaped rectangular appearance (

). It

should be noted that at least 24% of dogs with USMI have a normal
bladder neck position

[5,10,11]

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

Pressure proﬁlometry

Urethral pressure proﬁlometry has been used in research and clinical

settings to aid in diagnosis of USMI

. Urodynamic studies can reveal

reduced urethral resistance with subnormal urethral pressures

, reduced

leak point pressure, and reduced maximal urethral closure pressures

[13–15]

Diﬀerential diagnosis

Other causes of urinary incontinence in immature animals include ureteral

ectopia, ureterocele, urachal diverticula, impervious urachus, dyssynergia,
and cystitis. In adults, cystitis, neoplasia, prostatic disease, neurologic
abnormalities, and renal disease need to be ruled out.

Treatment

Medical management

Medical management is the initial treatment in USMI cases. The a-

adrenergic drug phenylpropanolamine (1.5 mg/kg every 8 hours adminis-
tered per os) increases smooth muscle contractility, increases elasticity, and
increases sensitivity to catecholamines

[16,17]

. Results vary, but reported

success in the prevention of involuntary urination may be as high as 85% in
treated dogs

[2,5,18]

. Recently, the use of sustained-release phenylpropa-

nolamine with diphenylpyraline in 11 bitches previously refractory to
phenylpropanolamine was reported with favorable results

. Unresponsive

animals may be treated with estrogen or combined medical treatment
protocols; however, there remains the potential for aplastic anemia and other

Fig. 1. Lateral radiographic view of a double-contrast cystogram revealing a pelvic bladder.
Note that greater than 5% of the cranial-caudal dimension of the distended bladder lies within
the pelvic canal.

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

side eﬀects of reproductive hormones. The reader is referred to other sources
for additional descriptions of medical management

[20,21]

Surgical management
Colposuspension

. Colposuspension is a surgical technique that is used to

treat USMI by relocating the bladder neck from an intrapelvic to an intra-
abdominal location (

Technique

. The dog is placed in dorsal recumbency, and a caudal

celiotomy is performed. A Foley urethral catheter can be placed, and
a ﬁnger or Carmalt forceps are introduced closed into the vagina to aid in
abdominal localization. Minimal blunt dissection is used to localize the

Fig. 2. (A) Lateral diagram of a dog with a pelvic bladder. (B) Lateral diagram of the same dog
after a colposuspension procedure. Notice the cranial and ventral displacement of the bladder
(and the proximal urethra) after colposuspension. (C) Intraoperative diagram of a colposus-
pension. The ventrolateral wall of the vagina is being pulled cranially as a mattress suture is
being placed. (Courtesy of the University of Tennessee College of Veterinary Medicine,
copyright 2003; with permission.)

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

urethra and vagina, and ﬁngers are used to separate the fascia between the
urethra and pubis. The prepubic tendon is isolated, and the inguinal canals
are avoided. The ventrolateral wall of the vagina is grasped with Allis or
Babcock forceps on either side of the urethra, and the vagina is pulled
cranially (see

Fig. 2C

). Sutures of 0 polypropylene are passed caudal to the

prepubic tendon, a lateral large bite of the vagina is taken, and the sutures
are returned through the ipsilateral abdominal wall just cranial to the
prepubic tendon, forming a mattress. Two sutures are generally placed in
middle- to large-sized dogs on either side of the vagina. Before tying, sutures
are pulled tight to ensure the urethra is not unduly compressed against the
pubis. After the procedure is completed, it should still be possible to pass the
tip of the Carmalt forceps between the urethra and pubis

Complications reported with the colposuspension procedure are un-

common but include postoperative straining to urinate, inability to
urinate, tearing of the sutures from the vagina, and recurrence of urinary
incontinence

[3,10,15,22]

. Dysuria in the immediate postoperative period

may be secondary to vaginal stimulation or intrapelvic stimulation
causing suppression of the micturition reﬂex

. Dyssynergia responsive

to diazepam administration has been suggested as the most common
cause of postoperative dysuria and may be exaggerated by prior estrogen
administration

. Subsequently, estrogen therapy should be discontin-

ued at least a week before the procedure. Another complication occurs if
the urethra is compressed, especially if sutures are placed too close to the
urethra. Complete outﬂow obstruction (which is extremely rare) may
necessitate suture removal.

Urethropexy

. Recently, a technique of urethropexy was described by

White

in 100 bitches. In this procedure, a caudal celiotomy is made,

a suture is placed in the apex of the bladder, and cranial traction allows
the urethra to be visualized. A 2/0 or 0 polypropylene suture is passed
caudal to the prepubic tendon into the abdomen and transversely through
the muscular layers of the urethra while gentle traction is maintained on
the bladder stay suture. Care is taken to avoid the urethral lumen,
although this is not always possible. The suture is then exited caudal to
the contralateral prepubic tendon. This suture is held in a hemostat, and
a second suture is passed 3 to 5 mm cranial to the ﬁrst; the bladder
suture is released before tying both urethropexy sutures

. Complica-

tions with this procedure were seen in 21% of cases and included
increased frequency of urination, dysuria, and, rarely, complete obstruc-
tion. All cases responded spontaneously within 14 days, except the 3
cases of outﬂow obstruction, in which the sutures were removed

Miscellaneous techniques

. Arnold and his colleagues

[2,24]

have reported on

the use of endoscopic urethral injection of Teﬂon (polytetraﬂuoroethylene)
and glutaraldehyde cross-linked collagen in bitches similar to the technique

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

used in women. A needle is advanced submucosally via urethroscopy, and
the bulking material acts to obstruct the urethral lumen partially

Urinary incontinence recurred in approximately two thirds of cases with
Teﬂon but often responded to repeat injection

. The cure rate with bovine

dermal collagen improves to approximately 50% with the ﬁrst injection
series

A transpelvic sling procedure has been described with and without

colposuspension in an eﬀort to increase midurethral pressure

. The

authors propose that dogs with low urethral pressures have a diﬀerent
pathophysiology than animals with normal urethral pressure and intrapelvic
bladders. Results of this study were comparable to those in which
colposuspension alone was used. Complications associated with the surgery
included dysuria, stranguria, and ﬁstulation

Cystourethropexy has been described by Massat et al

and is similar to

the urethropexy procedure. In this study of 10 bitches, dysuria was a common
complication; however, all were able to urinate voluntarily. Results were
excellent in 2 of the dogs and considered good in 4 of the dogs

Urethral lengthening is a technique where two V-shaped ﬂaps are made in

the ventral bladder wall and the resultant defect is closed to decrease the
diameter of the bladder neck

. Excellent results have been described in

a series of eight cats and in one dog

Prognosis

In a summary of owner-assessed outcomes in 150 cases of colposuspen-

sion, Holt

reported approximately 56% of bitches cured with surgery

alone; an additional 40% were signiﬁcantly improved and 10% exhibited
refractory incontinence. Similar results were observed in a study of 60
bitches undergoing colposuspension, with 40% cured, 42% markedly
improved or totally continent with concurrent medication, and 18% failing
to respond

. This study included 50 animals in which previous medical

management had been unsuccessful. Long-term deterioration occurred in 4
animals.

A recent prospective study evaluating colposuspension in 23 spayed

incontinent female dogs incorporated preoperative and 2-month postoper-
ative urethral pressure proﬁlometry and leak point pressure testing

Additional client interviews regarding outcome were performed up to 1 year
after surgery. In this study, 55% of dogs were continent at the 2-month
recheck; however, 1 year after surgery, only 14% were continent and
another 33% were described as greatly improved. Further inspection of the
data showed that when surgical and medical therapy was combined, 38%
were considered continent and a further 43% greatly improved. Client
satisfaction with the colposuspension procedure was high at 86%

Rawlings et al

have shown that the leak point pressure test increases

in bitches that exhibit improved urinary control after surgery. Using a lateral

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

radiograph of a retrograde vaginourethrogram, they also found that bitches
having a more caudal entry point of the urethra into the vestibule are more
likely to respond to the colposuspension

White’s results with urethropexy compared favorably with colposuspen-

sion

; 56 of 100 bitches were cured by surgery, and 27 were greatly

improved. Long-term follow-up over 12, 24, and 36 months showed
unexplained deterioration with time in 8 bitches. Interestingly, two thirds
of dogs that were reoperated on because of deterioration or failure were
greatly improved after a second surgery, suggesting that repeat surgery may
be justiﬁed

Summary

Medical management of USMI is a lifelong commitment and is

reasonably successful in many dogs. Although surgical treatment of USMI
does not provide continence in all animals with incontinence, it may be
considered as a viable option in many patients, especially when attempted
medical management has failed. Improved continence can be expected in
most animals.

Ectopic ureters

Pathophysiology

Any condition in which a ureter terminates at a site other than the normal

position at the trigone of the bladder is deﬁned as an ectopic ureter. Ectopic
ureters are the most common cause of juvenile urinary incontinence in
female dogs. Incidence has been estimated at 0.016% of small animal
accessions to veterinary teaching hospitals

[28,29]

. Ureteral ectopia is

a congenital condition, but heritability has been suggested as a possible
etiology because of established breed associations with the disease.
Labrador Retrievers, Golden Retrievers, Fox Terriers, Siberian Huskies,
Newfoundlands, Bulldogs, and Poodles have been cited as high-risk breeds

[30–33]

. Teratogens and other causes of congenital defects may be

implicated. Embryologically, if the metanephric duct arises too far cranially
on the mesonephric duct, the ureters may not reach the bladder. Instead,
female dogs may have ureters that open into the bladder neck, urethra,
uterus, vagina, or vestibule

. Ureters in male dogs may open into the

bladder neck, urethra, vas deferens, or seminal vesicles

[34,35]

Clinical features

Ectopic ureters are classiﬁed into two main categories, intramural and

extramural. Intramural ectopia is the most common form in dogs and
occurs when one or both ureters enter the urinary bladder wall dorsally near

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

the trigone at or near the normal location without entering the bladder
lumen

[28,32,34,36]

. Instead of opening into the trigone, intramural ectopic

ureters course submucosally to a more distal location. Multiple fenestrations
(oriﬁces within a ureter) or troughs (dished mucosal drain boards extending
into the urethra) may be present, which can complicate diagnosis and
treatment

[32,33,36,37]

Extramural ectopia results when one or both ureters completely bypass

the urinary bladder to insert further distally into the urogenital system.
Although relatively uncommon in dogs, extramural ectopia may be more
prevalent in cats

. Concurrent abnormalities associated with both types

of ectopic ureters have been reported to occur in as many as 94% of cases
and may include absent, small, or irregular kidneys; hydronephrosis;
pyelonephritis; pelvic bladder; and hydroureter

Diagnosis

Most ureteral ectopia occurs in young female dogs. A history of

intermittent or continuous urinary incontinence since birth is usually the
primary presenting complaint. Other causes of incontinence are possible in
young dogs; for this reason, a thorough workup must be performed to make
the diagnosis. The results of physical examination are often within normal
limits. Urine scald or perivulvar dermatitis is a possible clinical sign. A
minimum clinicopathologic database should include a complete blood cell
count, blood chemistry proﬁle, urinalysis, and urine culture. Survey
radiography, excretory urography, urethrocystography, pneumocystogra-
phy, ultrasonography, CT, and urethral pressure proﬁlometry have all been
used in establishing a diagnosis. Transurethral cystoscopy for direct
visualization of the trigone and the urethra remains the most insightful
and reliable method for achieving a diagnosis. A deﬁnitive diagnosis is made
via surgery or postmortem examination.

Minimum database

Analysis of blood and urine may help to rule out other causes of

incontinence. As well, results may help to detect concurrent disease.
McLaughlin and Miller

reported a concurrent urinary tract infection

in 64% of cases with ectopic ureters. Because the presence of infection may
lead to increased postoperative complications, treatment and resolution of
infections should be the goal before the initiation of surgery.

Radiography

Survey radiographs should be taken as a baseline to evaluate the

abdomen before further radiographic workup. Positive-contrast urethro-
cystograms may reveal ectopia if ureteral reﬂux occurs. Disadvantages of
using urethrocystography include a negative ﬁnding if reﬂux does not occur
and the inability to evaluate the upper urinary tract.

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

Excretory urography is a useful noninvasive procedure for diagnosing

ureteral ectopia. Alteration of the normal J-shaped ureterovesicular angle to
a straight line has been highly correlated to the presence of ureteral ectopia
(

. As well, preoperative excretory urograms can give insight to

prognosis by allowing visualization of the upper urinary tract, including the
kidneys and the ureters (

Pneumocystography in combination with an excretory urogram may help

to improve visualization of the distal ureter. In a study by Mason et al

a combination of the two techniques was more accurate in distinguishing
intramural from extramural ectopic ureters compared with excretory
urography alone.

Ultrasonography

Ultrasonography has been reported to correlate well with contrast

radiography for detection of ectopic ureters

[41,42]

. Ultrasonography may

be more accurate than radiography in determining the site of ureteral
termination and concurrent urogenital pathologic ﬁndings

Fig. 3. Excretory urograms with pneumocystograms (oblique views). (A) Distal ureter and J-
shaped ureterovesicular angle in a normal dog. (B) Bilateral hydroureter and ureteral ectopia in
a dog with urinary incontinence. Note the dilated ureter dorsal to and extending beyond the
neck of the bladder.

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

Transurethral cystoscopy

In skilled hands, cystourethroscopy has become increasingly valuable in

diagnosing ureteral ectopia and other urogenital anomalies in veterinary
patients

[43,44]

. Cystourethroscopy is more reliable than radiography

because direct visualization of both ureteral oriﬁces is possible

. The

ability to detect ureteral troughs, multiple ureteral fenestrations, and other
concurrent vaginal or vestibular abnormalities can make transurethral

Fig. 4. Excretory urogram with pneumocystogram. Lateral (A) and ventrodorsal (B)
radiographic views revealing bilateral ureteral ectopia with marked ureteral ectasia and
hydronephrosis. Changes are more severe on the right.

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M.G. Hoelzler, D.A. Lidbetter / Vet Clin Small Anim 34 (2004) 1057–1073

cystoscopy an invaluable technique for accurate preoperative surgical
planning

[43,44]

In a recent study by Samii et al

[45]

, CT was used as a method of diagnosis in

dogs with ectopic ureters. Results showed better agreement between CT and
cystoscopy and surgery or necropsy ﬁndings with regard to the presence and
site of ureteral ectopia compared with other imaging techniques.

Urethral pressure proﬁlometry

Concurrent urethral incompetence is common in dogs with ectopic

ureters. Although pressure proﬁlometry cannot be used to diagnose ureteral
ectopia deﬁnitively, it may be useful as a prognostic indicator for continued
postoperative incontinence. In a report by Lane et al

, urodynamic

measurements predicted postsurgical outcomes consistent with actual out-
comes in 89% of dogs. Preoperative urodynamic assessment of dogs with
ectopic ureters can help to identify concurrent abnormalities, such as
reduced bladder capacity and USMI

Diﬀerential diagnosis

In a series of 221 cases reported by Holt

, nearly 50% of juvenile animals

with urinary incontinence suﬀered from ureteral ectopia. Other disorders,
such as USMI, upper or lower urinary tract infection, cystic calculi, neurologic
conditions, and vestibulovaginal abnormalities must be ruled out.

Treatment

Surgery is the deﬁnitive treatment for ureteral ectopia. Many procedures

have been described, and their use varies with type and location of the
ureteral abnormality.

Ureteroneocystostomy

Ureteroneocystostomy, also known as ureteral reimplantation or ureter-

ovesicular anastomosis, is a technique used to correct extramural ectopic
ureters. A routine caudal midline celiotomy is performed. The caudal
abdomen is explored, and both ureters are identiﬁed. The aberrant ureter is
traced as caudally as possible before ligation and transsection. The distal 2
to 3 cm of the ureter is dissected away from surrounding periureteral fascia
with minimal collateral damage to vascular and neural structures. A stay
suture is applied through the distal end of the ureter. A ventral cystotomy is
made before performing a small stab incision through the dorsolateral
bladder wall. The stab mark is located, and small forceps are inserted and
tunneled between the urothelium and detrusor muscle toward the apex. The

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length of the tunnel should be approximately three times the normal ureteral
diameter to help minimize ureteral reﬂux

[48]

. Nevertheless, it is important

not to make the tunnel too long to avoid postoperative ureteral dilation and
hydronephrosis. The stay suture is then grasped with tunneled forceps to
draw the ureter through the tunnel and into the bladder. Once positioned
inside the bladder, the terminal ureter is excised. The remaining distal ureter
is spatulated with a short longitudinal incision (0.5–0.75 cm) and sutured to
the vesicular mucosa with four to six simple interrupted absorbable sutures
(

). Meticulous technique must be employed, and the use of magnifying

loupes is recommended to enhance visualization. Excessive trauma or
placement of too many sutures can lead to mucosal edema and scarring,
which could occlude the ureteral oriﬁce after surgery.

Other ureteroneocystostomy techniques, including implantation through

a transverse tunnel or an extravesicular ‘‘drop-in’’ technique, have been
described but have been shown to have increased rates of morbidity

[48–50]

Neoureterostomy

Because most ectopic ureters in dogs have an intramural morphology,

ureteroneocystostomy is usually not required. A conventional approach has
been to ligate the distal portion of the intramural ureter after creating
a proximal neoureterostomy. A new approach has been suggested by
McLoughlin and Chew

. With this technique, a routine caudal midline

celiotomy and ventral cystotomy are initially performed. The cystotomy

Fig. 5. Intraoperative diagram of an ureteroneocystostomy. (A) A dilated extramural ectopic
ureter has been transected, and a cystotomy has been performed. Hemostats are being used to
create a short tunnel through which the ureter will be introduced into the bladder. (B) The distal
ureter is spatulated and sutured to the vesicular mucosa with simple interrupted sutures. (C), A
newly created ureteral oriﬁce after ureteral reimplantation. (Courtesy of the University of
Tennessee College of Veterinary Medicine, copyright 2003; with permission.)

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incision is made far enough caudal to allow visualization of the aberrant
ureteral stoma distal to the trigone. In most instances, the incision must be
extended into the proximal urethra. Once the ectopics are identiﬁed, ureteral
troughs, fenestrations, and distal remnants are located. The intramural
ureteral remnant is then catheterized in retrograde fashion from the distally
displaced oriﬁce proximally. Using a combination of blunt and sharp
dissection, the ureteral remnant is excised from the urethra/distal trigone
with the help of the preplaced catheter. The distal ureteral remnant is
excised, and a stoma is fashioned with four to six interrupted mucosal
apposition sutures. Defects are closed with continuous suture patterns
(

Neoureterostomy takes advantage of a submucosal tunnel already

present at the vesicoureteral valve, thereby minimizing postoperative
valvular stenosis. Trauma to local vessels and nerves is less common.
Problems with neoureterostomy include continued incontinence if the distal
ureteral remnants and troughs are not completely resected. Visualization of
the proximal urethra in male dogs with prostatomegaly can be diﬃcult, and
catheterization of ectopic ureters in extremely small dogs may be impossible.
Therefore, ureteroneocystostomy may need to be considered in lieu of
neoureterostomy in these patients.

Nephroureterectomy

Complete excision of the aﬀected kidney and ectopic ureter is a salvage

procedure in the treatment of ureteral ectopia. The technique should only be

Fig. 6. Intraoperative diagram of a neoureterostomy. (A) An intramural ureteral remnant has
been catheterized and excised from the urethra and distal trigone. (B) The ureteral remnant has
been excised, and a stoma has been fashioned. The trigonal/urethral defect has been closed with
a continuous suture pattern. (Courtesy of the University of Tennessee College of Veterinary
Medicine, copyright 2003; with permission.)

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performed when severe pathologic ﬁndings are present and when the
contralateral kidney/ureter is functioning normally. Additionally, the fact
that many instances of ectopia are bilateral would warrant close inspection
of the entire urogenital system before contemplating nephroureterectomy

[39,41]

Complications

Complications of surgery vary with the method of repair and surgical

technique. Complications of ureteroneocystostomy include hydroureter,
hydronephrosis, cystitis, transient stenosis, anastomotic dehiscence, contin-
ued dysuria, and loss of normal ureteric peristalsis

[29,32,36,51]

. Compli-

cations of neoureterostomy include continued dysuria, cystitis, and potential
reﬂex dyssynergia

[29,32,51]

. Recanalization is a possible cause of post-

operative incontinence if the distal ureter is not completely resected

[40,52]

Surgeons must pay close attention to the possibility of multiple fenestrations
and the presence of ureteral troughs if postoperative incontinence is to be
minimized

Chronic urinary tract infections may lead to postoperative stricture,

hydroureter, pyelonephritis, and possible renal failure.

Prognosis

Of patients with ureteral ectopia, 44% to 67% have been reported to

have postoperative urinary incontinence

[33,36,40,46,51]

. Although residual

incontinence can occur after incomplete intramural ureteral remnant or
ureteral trough resection, incontinence has also been reported after ureteral
reimplantation and nephroureterectomy