Pyothorax in Cats: A Review - VCA Specialty Animal Hospitals

Volume 1, Issue 4
j
March/April 2011
Pyothorax in Cats: A Review
Valérie Sauvé, DVM, DACVECC
Fifth Avenue Veterinary Specialists
P
yothorax is defined as an accumulation
of purulent material in the pleural
space. Infection may be introduced
from the chest wall, diaphragm, lung
parenchyma, esophagus, or airways. The
most common cause of feline pyothorax
has long been thought to be bite wounds
to the chest from other cats, but recent
evidence points to another major cause: the
transmission of infection from the lungs.1
Other reported causes include aberrant
migration of Cuterebra, penetrating foreign
body, hematogenous or lymphatic diffusion
of distant infection, esophageal or tracheal
perforation, pulmonary abscess, lung
parasites, discospondylitis, neoplasia with
abscess, and iatrogenic infection.1-3
Orthogonal thoracic radiographs should be
obtained to confirm pleural effusion and
to determine if the fluid accumulation is
bilateral or unilateral. Radiographs may also
reveal underlying conditions such as a mass
or abscess and should be carefully reviewed
for evidence supporting other differential
diagnoses. Diagnostic or therapeutic
thoracocentesis (usually while the patient
is under sedation) is recommended. Any
effusion of unknown cause in the chest
should be sampled for fluid analysis and
cytology unless congestive heart failure is
suspected. If this is the case and there is
only a small amount of effusion, a diagnostic
thoracocentesis may be delayed pending
response to treatment with a diuretic.
Cats with pyothorax often present with
general signs of malaise such as lethargy and
decreased appetite. Tachypnea, coughing,
and weight loss may be reported and some
animals will be dyspneic. Symptoms such as
hypersalivation and bradycardia have been
associated with a worse outcome.2 On physical
examination, the majority of patients will be
febrile. Auscultation may reveal ventrally
quiet lung sounds, unilaterally or bilaterally.
Wounds may be present over the chest but
this is a rare finding. A complete, attentive
physical examination is recommended as
multisystemic findings may help direct the
diagnostic and therapeutic approach.
Patients with pyothorax will have an exudative
effusion featuring both a high WBC count and
total solids (>3.0 g/dL). Cytology of the effusion
can be performed in house but should also be
submitted to a clinical pathologist for review, as
filamentous bacteria and neoplastic cells may
be more difficult to identify. The cytology of
pyothorax fluid will typically show degenerative
neutrophils and intracellular and/or extracellular
bacteria. Ideally, cultures should be submitted,
both aerobic and anaerobic, on all patients
before the commencement of antibiotic therapy.
Anaerobes with or without aerobic bacteria are
most commonly involved. Acid fast testing on
bacterial isolate may be helpful if filamentous
bacteria are present in the sample. If positive,
TMS should be added to the antibiotics.
Blood tests and advanced imaging techniques
can be used to investigate the cause of
the pyothorax and to help the clinician in
discussing prognosis with the owners. Baseline
testing including CBC, biochemistry, and
FeLV/FIV is recommended. Patients with
higher WBC counts have been found to have
a better outcome.2 FIP PCR of the effusion
may be indicated. Toxoplasmosis IgG/IgM
and Cryptococcus AG may be performed
if parenchymal lung disease is present.
Abdominal ultrasound can be used to look for
other organ system involvement and thoracic
ultrasound may identify masses, abscesses,
or pocketed fluid accumulation. Ultrasound
is also very useful in guiding thoracocentesis
in patients with a small amount of effusion.
Thoracic computed tomography (CT scan)
may become more commonly performed in
the future to determine the underlying disease
and need for surgical exploration.
Treatment is mostly focused on draining
the infection. Repeated thoracocenteses are
associated with poor outcome and cannot be
recommended. Unilateral but usually bilateral
chest tubes should be placed as soon as the
patient is stable. Performing a therapeutic
thoracocentesis and administering IV fluids
and oxygen prior to general anesthesia is
Continued on page 2
Pyothorax in Cats
Continued from page 1
beneficial to make the patient more stable
for placement. Patients with thoracostomy
tubes need to be monitored 24 hours a day
because disconnection can cause a severe
pneumothorax. If the fluid is thick, flushing
the tubes with sterile saline may be beneficial
as this will help to dislodge the purulent
material. Approximately 10 mL/kg of sterile
warm NaCl 0.9% is slowly infused, the animal
is turned and rotated and then the fluid is
removed. This can be performed 2-4 times
per day. It is important to note the amount
of fluid flushed and retrieved as the inflamed
pleura may reabsorb fluid and this needs to be
accounted for in the IV fluids plan.
Case Review: Luna
Luna is a 12 year old DSH who presented to FAVS’
cardiology service for evaluation of pleural effusion.
She had a history of weight loss, inappetence and
lethargy for several weeks. Her blood work at Gramercy
Park Animal Hospital showed severe neutrophilia
with a left shift. She received enrofloxacin for two
Luna with her bilateral
weeks which improved her clinical signs, but her
thoracostomy tubes.
neutrophilia was persistent and she began to decline
once the antibiotics were discontinued. She had also developed vomiting and mild dyspnea.
An echocardiogram showed Luna’s cardiac structure and function
to be normal. Our cardiologist, Dr. Sophy Jesty, performed a
diagnostic thoracentesis under sedation which revealed a
septic suppurative exudate. Luna was then transferred to the
Critical Care service. She received intravenous fluids, including
Hetastarch, to address her hypotension and dehydration
overnight, and started antibiotic therapy. Her clotting times were
prolonged therefore she received fresh frozen plasma and vitamin
K1. Her oxygen saturation was within normal limits so she did
not require oxygen supplementation. Luna’s CBC showed that the
neutrophilia and left shift had worsened significantly but she was
afebrile. Bacterial cultures of the pleural effusion were negative
although the fluid analysis and cytology were confirmatory for
pyothorax. The next day bilateral thoracostomy tubes
were placed with Luna under general anesthesia.
Supportive care is also essential for the
recovery of these patients. Intravenous
fluids are needed as well as parenteral pain
medication. A fentanyl/ketamine CRI is
often used for analgesia as the chest tubes
are painful. Intravenous antibiotics are also
indicated. Broad spectrum antibiotics are used
while awaiting culture results, and anaerobic
coverage is mandatory. The combination
of enrofloxacin and ampicillin-sulbactam
is usually the first line. Respiratory support
such as bronchodilators, nebulization, and
oxygen therapy may also be needed, and
electrolytes and nutritional intake should be
carefully monitored.
Luna remained in the hospital for 7 days, receiving
IV enrofloxacin, ampicillin-sulbactam, famotidine,
and a ketamine/fentanyl CRI. She also benefited from
nasogastric Clinicare feedings. The tubes were initially
flushed 3 times per day with 10 ml/kg of warm sterile
saline. The left-sided tube was removed after 4 days
as it was no longer productive and the left lung was
Lateral and v/d images showing Luna’s completely normal on
pleural effusion and chest tube placement. follow-up radiographs.
The right-sided tube was removed after 7 days. The last radiographs of
her stay showed persistent right sided infiltrates. She was discharged on
amoxicillin-clavulanic acid and enrofloxacin orally.
The majority of patients will require an
average of 4-6 days of treatment before the
chest tubes can be removed. Exploratory
thoracotomy or thoracoscopy may benefit
some patients with compartmentalized fluid
or a poor response to medical treatment,
and early surgical intervention is indicated
in those patients with a thoracic wall
lesion, intrathoracic mass, foreign body, or
perforated esophagus. In human patients,
biochemical values such as glucose, pH, and
lactate are used to determine the need for
invasive treatment, however this has not been
studied in animals. A CT scan may be useful
in planning a surgical approach. Antibiotic
therapy should be continued for 4-6 weeks,
ending 1-2 weeks after complete resolution
of radiographic changes.
The prognosis for cats with pyothorax
treated as inpatients with thoracostomy tubes
is fair to good, with a favorable outcome
reported in 66-95% of cases.1-3 Thoracotomy
was required only in a minority of pets in
published studies, and the need for surgery
was not indicative of a worse prognosis. j
2
At the first recheck, Luna unfortunately had persistent unilateral pleural
effusion. Thoracocentesis revealed a low- to moderate-grade chronic-active
suppurative inflammation. FIP PCR was negative. However, her WBC count was
improving and she was doing very well at home, gaining weight and having a
good appetite. The owners declined CT and exploratory thoracotomy and elected
to continue oral antibiotics. Luna was then rechecked at Gramercy Park Animal
Hospital by Dr. Karen Feibusch and her radiographs and CBC normalized after
8 weeks of antibiotics. Four months after her stay at our hospital, she is doing
wonderfully at home!
Luna at home a few months after her stay!
References and recommended reading
Barrs VR. Feline pyothorax: a retrospective study of 27 cases in Australia. J Feline Med Surg
2005;7(4):211-222.
2
Waddell LS et al. Risk factors, prognostic indicators, and outcome of pyothorax in cats: 80
cases (1986-1999). J Am Vet Med Assoc 2002;221(6):819-824.
3
MacPhail CM. Medical and Surgical Management of Pyothorax. Vet Clin North Am Small
Anim Pract. 2007;37(5):975-988, viii.
1
Full Circle Forum
Case Report: Ivermectin Toxicity and
Treatment with Lipid Emulsions
Melissa L. Holahan, DVM, DACVECC
VCA Shoreline Veterinary Referral & Emergency Center
“D
usty,” a 3 year old, 33 kg, intact
male Labrador Retriever was
presented for evaluation of
suspected ivermectin toxicity. He was seen
ingesting approximately 600 mg of an equine
ivermectin 1.87% paste 12 hours prior to
presentation (maximum dose of 18 mg/kg
ivermectin). Initial physical exam revealed
bradycardia (HR=70 bpm), blindness, mild
tremors, vocalization, hypersalivation and
severe pelvic limb ataxia. The dog was
disoriented, agitated, and running into walls.
Further neurologic examination revealed
mydriasis, absent menace, and absent pupillary
light reflexes (direct and consensual) bilaterally,
along with conscious proprioceptive deficits in
the pelvic limbs. Dusty was unable to navigate
a maze or track a cotton ball.
Diagnostic tests included a serum chemistry
and complete blood count which were both
unremarkable. Gastric emptying procedures were
not pursued due to the time delay between ingestion
and admission (12 hours). Anecdotal evidence
of successful treatment of canine ivermectin
toxicosis with the novel therapy of intravenous
lipid emulsion (ILE) administration has been
documented. Based on this evidence we elected
to treat the dog with intravenous fat emulsion
(Liposyn II 20%) one hour post admission.
Dusty was given a 1.5 mL/kg intravenous bolus
of Liposyn followed by a constant rate infusion
of 0.25 mL/kg/min for sixty minutes.
Post-lipid emulsion therapy, Dusty’s pupillary
light reflexes were markedly improved and he
was able to navigate a maze and track cotton
balls. He also had mild improvement in the
pelvic limb ataxia. The dog was also treated
with 1g/kg of activated charcoal (Toxiban with
Sorbitol) orally. He was discharged 6 hours
post admission due to financial constraints and
was sent home with three additional doses of
activated charcoal without sorbitol to be given
every 6 hours. Dusty was reported by the family
to be clinically normal 12 hours after initial
presentation (24 hours post ingestion).
A toxicology panel was submitted for macrolide
endectocides and reported a positive serum
ivermectin level of 2540 ppb prior to lipid
infusion. Immediately following lipid infusion,
March/April 2011
the ivermectin level was less than 30 ppb. This
case report is the first to document ivermectin
levels pre- and post-lipid emulsion therapy. These
results provide further evidence that lipid emulsion
therapy may be valuable in the initial treatment
of ivermectin toxicity. Its use is also supported by
recent case studies involving treatment of various
toxicities including avermectins.1,2
patient’s exposure is unknown, other toxins such
as carbamate and organophosphate insecticides,
tremorogenic mycotoxins, sedatives or muscle
relaxants should be considered. A macrolide
endectocides toxicology panel can be requested
through the California Animal Health &
Food Safety Laboratory System to test for
avermectins if suspected.
Ivermectin is a broad-spectrum antiparasitic drug
commonly used against nematode infestations
and external parasites. Ivermectin is licensed for
use in cats and dogs as heartworm prevention.
Dogs can develop clinical signs of ivermectin
toxicity at doses >2.5 mg/kg from accidental use
or ingestion of equine or livestock deworming
products. Collies and other herding dogs are
more sensitive to ivermectin toxicity due to
potential deficiencies in one of the MDR1-type
P-glycoproteins.3 Toxicity has also been reported
in other breeds and cats at approved labeled doses.
Other avermectins that have similar mechanisms
of action and toxicity include moxidectin,
eprinomectin, selamectin, and doramectin.
Gastric decontamination is the first step in
treatment unless contraindicated (lack of gag,
decreased mentation); follow-up therapy is
largely supportive. Repeated doses of activated
charcoal should be administered due to
enterohepatic recirculation. Those patients that
are symptomatic may require anticonvulsant
therapy. Diazepam and other benzodiazepines
are believed to potentiate the effects of ivermectin
and prolong recovery due to their GABAenhancing properties.5,6 Alternative drugs for
anticonvulsant therapy include phenobarbital
and propofol. Patients that are stuporous or
comatose should be monitored closely to ensure
that they have a gag reflex and can maintain
their airway. Those patients that have lost their
gag reflex or become hypercapnic may require
endotracheal intubation and ventilator support.
Ivermectin is an agonist of invertebratespecific,
glutamate-activated,
inhibitory
chloride channels causing flaccid paralysis and
subsequent death in nematodes and mites. The
toxic effects seen in mammals after ivermectin
ingestion are due to the similarity of the
invertebrate-specific channels to the vertebrate
γ-aminobutyric acid (GABAA)-gated channels
that inhibit interneurons in the central nervous
system. At therapeutic doses, the blood-brain
barrier protects against these effects. At high
doses, ivermectin potentiates these channels
causing hyperpolarization of cell membranes,
thus preventing neuronal depolarization.4
Clinical signs reported with ivermectin toxicity
include ataxia, vocalization, disorientation,
hyperesthesia, blindness, weakness, mydriasis,
and bradycardia. Severe cases may develop
seizures, generalized weakness, respiratory
depression, stupor or coma. Signs may manifest
as early as 2-4 hours post-ingestion but may
be delayed up to 24 hours. These signs can
also progress for 5-7 days post-exposure and
patients should be observed closely during this
time. Diagnosis is typically based on witnessed
or suspected ingestion of ivermectin. If the
Anecdotal evidence has suggested that ILEs may
be a promising antidote for ivermectin toxicity.7
ILEs such as Liposyn II (20% soybean oil, 1.2%
egg phosphatides and 2.5% glycerin in water for
injection) are traditionally used as a component of
parenteral nutrition therapy and have a 1 year shelf
life. Possible acute risks of short-term ILE therapy
include thrombophlebitis during peripheral IV
administration, anaphylaxis, and fat emboli.
However, ILEs have a very safe track record
based on their frequent and longstanding use in
parenteral nutrition therapy in critically ill patients.
One proposed mechanism of lipid therapy is
that the exogenous lipid provides an alternative
source for binding of lipid soluble drugs,
including local anesthetics and avermectins.8
This “lipid sink” theory holds that lipophilic
drug molecules (such as ivermectin) shift into
a lipemic plasma compartment making them
unavailable to the tissue. Other proposed
mechanisms including increased metabolism,
distribution, or partition of drug molecules away
from receptors into lipids within the tissues.8
Continued on page 4
3
Ivermectin Toxicity & Treatment w/ Lipid Emulsions
Lipid emulsions may also have a role in the
treatment of permethrin toxicosis due its high
lipid partition coefficient but currently no cases
have been reported in veterinary medicine.
Dosages of intravenous lipid emulsions (such
as 20% Intralipid) are extrapolated from
human medicine: 1.5 mL/kg as an initial
bolus, followed by 0.25 mL/kg/min for 3060 minutes. Boluses could be repeated 1-2
times with a maximum total dose of 8 mL/
kg recommended. Although standards for the
optimal and maximal doses to be given, the
appropriate rate of administration, and the
duration of therapy have not been established
in veterinary medicine, the anecdotal evidence
of using lipid emulsion therapy early in
ivermectin toxicity is encouraging.
In summary, intravenous lipid therapy is a novel
Continued from page 3
treatment approach for ivermectin toxicity. Its use
is supported by recent research and case studies
involving lipid administration for bupivacaine8
and other fat-soluble toxins. Lipid administration
in this case appeared to reverse the dog’s clinical
signs and markedly decreased the ivermectin level
present in the serum. This therapy may also prove
to be beneficial with other fat-soluble toxins, such
as beta-blockers and permethrins. j
References:
Crandell DE, Weinberg GL. Moxidectin toxicosis in a puppy successfully treated with intravenous lipids. J of Vet Crit Care. 2009;19:181-186.
O’Brien TQ, Clark-Price SC, Evans EE, DiFazio R, McMichael MA. Infusion of a lipid emulsion to treat lidocaine intoxication in a cat. J Am Vet Med
Assoc. 2010;237(12):1455-1458.
3
Mealey KL, Bentjen SA, Gay JM, Cantor GH. Ivermectin sensitivity in Collies is associated with a deletion mutation of the MDR1 gene. Pharmacogenetics. 2001;11(8):727–733.
4
Peterson ME, Talcott PA. Small Animal Toxicology, 2nd ed. Philadelphia:WB Saunders, 2006.
5
Williams M, Risley EA. Ivermectin interactions with benzodiazepine receptors in rat cortex and cerebellum in vitro. J Neurochem. 1984;42(3):745–753.
6
Williams M, Yarbrough GG. Enhancement of in vitro binding and some of the pharmacological properties of diazepam by a novel anthelmintic agent,
avermectin B1a. Eur J Pharmacol. 1979;56(3):273–276.
7
LipidRescue Web site. Available at: www.lipidrescue.org. Accessed January 24, 2011.
8
Weinberg G, Ripper R, Feinstein DL, Hoffman W. Lipid emulsion infusion rescues dogs from bupivacaine-induced cardiac toxicity. Reg Anesth Pain
Med. 2003;28(3):198-202.
1
2
Referring Veterinarians As Partners
Dermatophytosis, Drug Resistant Superficial
Pyoderma, and Atopic Dermatitis in a Dog
Jeanne Budgin, DVM, DACVD, Animal Specialty Center, and Patricia Doherty, DVM
“S
ophie,” a 7 year old spayed female
Vizsla, initially presented to Dr. Patricia
Doherty with a four year history of
pruritic, non-seasonal skin disease. A previous
blood allergy panel (HESKA Allercept) had
been performed and revealed positive reactions
to house dust mites, storage mites, several molds,
and one tree. Allergen specific immunotherapy
(ASIT) had been administered for a two
year period without improvement and was
subsequently discontinued. The skin condition
had improved with prednisone therapy in the
past, however unacceptable side effects including
marked polyuria, polydipsia, and polyphagia were
reported. Dr. Doherty had recently prescribed
oral Atopica at 5.3 mg/kg every 24 hours and
recommended a food elimination dietary trial
with Hill’s z/d Ultra to evaluate for an adverse
reaction to food. One month after initiating
treatment with Atopica, new, mildly pruritic skin
lesions consisting of multifocal annular patches of
alopecia, erythema, and scale were present on the
convex pinnae and paralumbar area. Dr. Doherty
performed a deep skin scraping (negative),
4
collected a fungal (DTM) culture, and initiated
oral antibiotic therapy (cephalexin, 26 mg/kg
every 12 hours x 21 days). After two weeks, the
fungal culture was positive for Microsporum canis.
The Atopica was lowered to 5.3 mg/kg every 48
hours; twice weekly KetoChlor shampoo and
twice daily 1% miconazole lotion were prescribed.
Though the skin lesions initially improved,
Sophie continued to develop new lesions and was
referred to the Dermatology and Allergy Service
at Animal Specialty Center.
Our dermatological evaluation yielded several
findings. There were patches of alopecia with
adherent white scale and increased epilation near
the right medial canthus, surrounding the base of
each ear, and covering the convex aspects of both
pinnae. There were multiple papules and heavy
yellow crust on the concave aspect of the right
pinna, and the lateral aspects of the muzzle, dorsal
head, medial left forelimb, lateral thorax, and
flanks exhibited mutifocal papules and plaques
with fine surface scale. Skin cytology via acetate
tape preparation revealed no significant findings
and flea combing was negative. Wood’s lamp
examination was positive in multiple sites. Sophie’s
assessment was generalized dermatophytosis
due to M. canis. The history of glucocorticoidresponsive dermatosis, as well as the results of the
previous blood allergy panel, supported atopic
dermatitis but due to the non-seasonal nature
of disease, an adverse reaction to food could not
be ruled out. The treatment plan included oral
itraconazole at 5.3 mg/kg every 24 hours, lime
sulfur dips every five days, and discontinuation of
Atopica. The food elimination dietary trial would
be continued for 10-12 weeks with Interceptor
being replaced by Revolution to eliminate
flavored medication. The importance of
confinement and environmental treatment
to destroy infective spores were emphasized.
Blood work would be performed monthly
with Dr. Doherty to screen for itraconazoleassociated hepatotoxicity.
At re-examination four weeks later, the skin
lesions had improved with treatment, however
pruritus was more generalized. Additional
skin lesions included the presence of multiple
papules and pustules on the ventral abdomen.
Full Circle Forum
Skin cytology (impression smear) revealed
moderate neutrophils (often degenerate) with
intracellular and extracellular cocci bacteria,
consistent with superficial pyoderma.
A fungal culture was repeated (negative) and
oral Simplicef at 7.8 mg/kg every 24 hours was
prescribed. Douxo Calm shampoo was also
recommended 1-2 times weekly (prior to lime
sulfur dip) to alleviate pruritus. The itraconazole
was continued with a one-week-on, oneweek-off regime, as this drug accumulates
and maintains adequate concentrations in
keratinized tissue to allow for pulse dosing.
Upon re-examination four weeks later, the skin
lesions were resolving, however pruritus was
escalating. There was still evidence of superficial
pyoderma on skin cytology. Because both
bacterial infection and resolving dermatophytosis
were present and may contribute to pruritus, it
was difficult to determine if improvement was
evident on the dietary trial. However, because
atopy is far more common that adverse reaction
to food, an intradermal skin test (IDST), an
aerobic culture to evaluate for drug resistant
pyoderma, and a fungal culture were performed.
The IDST revealed strong positive reactions to
house dust mites, storage mites, and cockroaches,
as well as several grasses, weeds, and trees. Aerobic
culture revealed methicillin resistant Staphylococcus
pseudintermedius sensitive only to amikacin and
rifampin. ASIT was re-initiated based on the
combined results of both skin and blood allergy
tests. Intensive topical treatments were instituted
including daily bathing in a Douxo Chlorhexidine
PS shampoo followed by the application of 2%
mupirocin ointment.
Sophie’s skin infections resolved and she was
maintained on a low dose of Temaril-P to
relieve pruritus associated with allergic skin
disease and continued receiving allergen specific
immunotherapy and weekly shampoo therapy.
A structured food challenge was performed and
no exacerbation in pruritus was appreciated, so
food allergy was considered unlikely.
The complexity and outcome of this case
demonstrates several important considerations
when managing patients with dermatologic
disease. While dermatophytosis is relatively
uncommon in adult dogs, and it is difficult to
determine if previous drug therapy (prednisone
and cyclosporine) predisposed this patient
to infection, it is important to include as a
differential for folliculitis, especially when
skin scrapings are negative and the response
to antibiotic treatment is poor. Environmental
treatment and oral antifungal therapy are both
important for effective management.
An aerobic culture is indicated when active
cytology persists after an appropriate dose and
duration of antibiotic therapy. Multi-drug
resistant pyoderma is on the rise in veterinary
medicine and antibiotic therapy must be prescribed
judiciously. In many cases, topical therapy is the
only option, and while often effective, requires a
very dedicated owner. Identifying and managing
the underlying cause of pyoderma, especially
when recurrent, is essential.
Finally, ASIT is most effective when based on an
IDST, or a combination of intradermal skin and
blood allergy testing, and often allows for less
dependence on medications such as glucocorticoids
and cyclosporine, which was evident in this case.
To date, Sophie continues to do well with
no clinical signs of skin disease. Her atopic
dermatitis is well controlled and she is reexamined every six months to monitor for any
signs of infection or modification of therapy.j
Elbow Dysplasia in the Dog
Matthew Palmisano, DVM, Diplomate ACVS
VCA Veterinary Referral & Emergency Center
“C
ody,” a 9 month old, male castrated
Labrador Retriever, presented to
VCA VREC with a three month
history of intermittent bilateral forelimb lameness.
The lameness improved with rest but worsened
with any activity. He was otherwise normal.
Orthopedic evaluation revealed shifting forelimb
lameness. Both forelimbs had slight abduction
at the elbow, with palpable joint effusion. Both
elbows were painful on extension, and they were
particularly painful with palpation of the medial
compartment of the elbow joint. Other general
and orthopedic findings were unremarkable.
Radiographs of the elbows were taken (see figures
1-3). Radiographic findings include subtrochlear
bone sclerosis in the region of the medial coronoid
process in both elbow joints on the lateral views.
Cody’s signalment, physical examination, and
radiographic findings are most consistent with
elbow dysplasia, most specifically, fragmented
medial coronoid process (FMCP) disease.
March/April 2011
Elbow dysplasia is a term that encompasses four
disease processes: FMCP, ununited anconeal
process (UAP), osteochondritis dissecans (OCD)
of the medial aspect of the humeral condyle,
and asynchronous radius-ulna growth. While
considered separate, these diseases often occur
together and may be interrelated. For example,
both FMCP and UAP are thought to be caused
by asynchronous growth between the radius and
ulna. Also, a percentage of dogs with FMCP
have OCD on the opposing articular surface
(kissing lesions). In this region of the country,
FMCP is the most common disease condition
associated with the elbow in the young, growing
large-breed dog.
Diagnosis of FMCP is primarily made based
on signalment, PE, and radiography. Lateral
radiographic views of the elbow joints are most
helpful for diagnosis of FMCP. Remodeling of the
subchondral bone beneath the FMCP causes the
subtrochlear sclerosis seen on the lateral views. AP
views are not often helpful, and oblique views of the
elbow are also relatively insensitive at identifying
the fragment. Other imaging modalities include
CT imaging and MRI. Both techniques have
varying sensitivity and specificity based on the
literature. Direct visualization of the fragment via
arthroscopy can provide a definitive diagnosis.
Treatment options include medical therapy, open
arthrotomy, and arthroscopy. Medical therapy
includes exercise restriction, chondroprotection,
and NSAID therapy. Open arthrotomy involves
making a 1-2cm incision on the medial aspect
of the elbow joint and removing the fragment
under direct visualization. Studies that have
compared the long-term outcome of medical
therapy to arthrotomy did not find any
significant difference in clinical improvement
between the two treatments.1,2 Many patients in
both treatment groups did not improve.
Minimally invasive surgery has gained popularity
over the past decade. Arthroscopic surgery of
Continued on page 6
5
Elbow Dysplasia in the Dog
the elbow joint is one of the most commonly
performed minimally invasive procedures in
veterinary orthopedics. Studies comparing
arthroscopic FMCP removal to arthrotomy and
medical therapy found that arthroscopy was
superior to both open surgery and conservative
management for improvement in clinical signs.1,2
Arthroscopy’s improved success is likely due to
the decreased trauma to the joint, as well as more
efficient fragment removal and debridement
of the subchondral bone, due the fact that the
operative field is magnified and bloodless.3
Figures 1-3: Anteriorposterior (top), right
lateral (lower right),
and left lateral (lower
left) radiographs of the
elbow joints. Note the
subtrochlear sclerosis
(asterisks) in the
region of the medial
coronoid process, and
hourglassing (arrows)
of the trabecular bone of
the proximal ulna.
For Cody, we recommended single-session
arthroscopic FMCP removal from both elbow
joints. The fragment in each elbow was identified
at surgery and removed through a small
instrument portal. The remaining subchondral
bony bed was debrided down to healthy bleeding
bone. The exposed healthy bone surface forms
a fibrin clot, which will eventually remodel into
fibrocartilage. Cody recovered uneventfully from
the surgery and anesthesia.
Cody was discharged the following day with
instructions for short leash walks for six
weeks. Deramaxx, tramadol, and Clavamox
therapy were administered for one week postoperatively. In addition, Dasuquin therapy was
recommended long-term for joint support. Skin
sutures were removed at two weeks, and Cody
was rechecked at six weeks before returning to
normal activity. At that point, he was essentially
sound, so he was gradually returned to normal
activity over the following two weeks. j
Continued from page 5
References:
Bouck GR, Miller CW, Taves CL. A comparison of surgical and medical management of fragmented medial coronoid process and OCD of the canine elbow. Vet Comp Orthop Traumatol.
1995;8:177-183.
2
Evans RB, Gordon-Evans WJ, Conzemius MG. Comparison of three methods for the management of fragmented medial coronoid process in the dog. A systematic review and meta-analysis. Vet
Comp Orthop Traumatol. 2008;21:106-109.
3
Beale BS, et al. Arthroscopically-assisted surgery of the elbow joint. In: Small Animal Arthroscopy.
Philadelphia, Pa: W.B. Saunders Co.; 2003:51-79.
1
Continuous Renal Replacement Therapy:
Information for Clinicians
Melissa Holahan, DVM, DACVECC; Samuel Durkan, DVM, DACVECC;
Larry Berkwitt, DVM, DACVIM; Danna Torre, DVM, DACVECC
C
ontinuous
Renal
Replacement
Therapy (CRRT) is now available at
VCA Shoreline Veterinary Referral
& Emergency Center in Shelton, CT. The
information outlined here will help you
determine if you have a patient who may be a
candidate for this treatment.
CRRT is a form of hemodialysis used when
gradual toxin (especially BUN) or water
removal is indicated in critically ill patients.
6
The VCA Shoreline specialty team will
evaluate each patient on an individual basis
to determine if this therapy is appropriate.
In addition to CRRT, there are other forms
of dialysis available for companion animals:
• Intermittent hemodialysis is fast and
efficient at clearing small solutes such as
blood urea nitrogen (BUN), potassium,
and certain toxins, like antifreeze.
• Peritoneal dialysis removes toxins from
the body by way of fluid placed into the
abdomen.
When patients are first started on
intermittent hemodialysis, they must be
introduced gradually to allow them time to
adapt. Therefore, the first few treatments
tend to be shorter with slower blood flow.
The speed and time of the treatment is
gradually increased each day. Treatments are
typically performed daily for several days in
Full Circle Forum
a row until the patient has adapted. Once
the patient is adapted to the treatment, the
treatments generally occur three days a week,
until the patient recovers renal function.
The standard treatment time during this
maintenance phase is usually four hours for
cats and five hours for dogs.
are not going to heal. Most of the time,
any potential kidney repair that is going to
occur will happen in the first four weeks.
Occasionally the kidneys will heal sooner,
and other times they take longer than four
weeks to heal. There is no way to predict
recovery time at the outset of treatment.
When patients are started on CRRT, the
treatment is intended to be essentially
continuous
which
requires
24-hour
specialized nursing care. After these patients
are stable, they will (in most cases) be
switched to intermittent hemodialysis. VCA
Shoreline offers CRRT for dogs in acute
renal failure who are at least 10 kg (22 lbs).
In cases of chronic renal failure, the kidneys are
permanently damaged. Dialysis is continued
three times a week for the rest of the patient’s
life. In this case, kidney transplant is the only
alternative to chronic dialysis.
Which patients may benefit from
dialysis?
• Patients with acute kidney failure
• Patients who have not responded to
standard therapy within 24 hrs. (fluids,
medication, etc.)
• Patients with life-threatening complications
of kidney failure or its treatment (e.g. high
potassium levels or fluid within the lungs)
• Patients who have ingested toxins that can
be removed with dialysis
Dialysis replaces many of the functions of
the kidneys, but it cannot replace them all.
Therefore, dialysis patients in the initial
stages of treatment will need to stay in the
hospital between treatments for ongoing
medical care. This includes fluid treatments,
antibiotics, and anti-ulcer medications,
among others. Because patients with
advanced renal disease often suffer nausea
and vomiting, medications are typically
administered parenterally. Hospitalization is
also required to allow for the frequent patient
monitoring that is necessary. Monitoring
includes blood pressure, urine production,
blood counts, and serum chemistry.
Is home care needed?
Once the patient is stable and can take
medications by mouth or by a feeding tube
(which is frequently needed during the
recovery stage), they may be able to go home
between treatments. The patient would
then return on a treatment schedule of three
days per week. The catheter is covered by
a bandage and needs no care at home other
than to keep it clean and dry.
How long is dialysis continued?
When dialysis is used for acute renal failure,
it is continued until the kidneys recover
function or it becomes clear that the kidneys
March/April 2011
Can dialysis prolong life?
An animal with complete loss of kidney
function will not live more than four to five
days without treatment. However, it can take
up to four weeks for the kidneys to recover
from acute injuries. Dialysis is intended to
support patients during this healing time.
However, not all pets with acute kidney
failure can recover, even with dialysis. About
50% of patients who have the support of
dialysis will survive. Unfortunately, the
other 50% cannot be saved despite all of
our efforts. Of those who live, 50% recover
completely with no lasting effects while the
other 50% may end up with chronic kidney
disease that requires ongoing dietary and
medical management.
Some types of renal disease have a better
chance of recovery than others. Damage
caused by infections and ischemia to the
kidney can be successfully treated 50-80%
of the time, whereas the chance of recovery
from toxin induced kidney failure is only
20% with dialysis.
Pets with chronic kidney failure on lifelong
dialysis may live a year longer than they
would have without dialysis.
How will animals feel during dialysis?
Most animals tolerate dialysis well. After the
initial dialysis treatment they may seem to
look their worst, but they usually show signs
of feeling better within 24-72 hours. They
often appear much more alert, they show
interest in being petted, and cats will begin
grooming themselves. Some patients will
even begin to eat and drink when often they
could not be coaxed to do so before dialysis.
During the treatment, patients sit in a quiet
room on a cushioned table with a blanket. A
technician is always by their side to monitor
their vital signs and respond to potential
complications.
Animals don’t seem to
experience the same degree of exhaustion
after dialysis that many people experience.
What are the risks of dialysis?
Any major medical or surgical intervention
carries risks, and this also is true for dialysis.
Some of the risks are due to the procedure
and some are due to the kidney failure itself.
The staff is trained to identify potential
problems so immediate treatment or
preventative measures can be initiated.
How much does dialysis cost?
Hemodialysis is an intensive treatment that
requires sophisticated equipment and 24
hour care with specially trained staff. Once
the pet has been evaluated and a therapeutic
plan which includes dialysis is in place,
clients will be given an estimate for the first
week of treatment.
Are there alternatives to dialysis?
Almost all patients considered to be
candidates for dialysis have already failed
to respond to medical management, and
the only other alternative is euthanasia. In
some cases, particularly cats with chronic
kidney disease, kidney transplantation may
be available and would usually be preferred
over dialysis. Kidney transplantation is not
suitable for cats that are severely ill, and
sometimes dialysis is necessary for a few
weeks to allow time to arrange a kidney
transplant. If kidney obstruction (typically
from kidney stones or scarring after kidney
stones) is the cause of the acute kidney
failure, sometimes dialysis can be avoided
by placing a tube directly into the kidney to
drain the urine.
How to have your patients evaluated
for dialysis at VCA Shoreline:
If you have a patient in whom you have
diagnosed kidney failure, and you and the
owner are interested in pursing hemodialysis,
please contact a member of the team to
discuss your pet’s case at (203) 929-8600.
Dialysis is offered by referral only and must
be approved by one of our team members.
If your patient needs emergency dialysis
after hours, please instruct your client to
bring them directly to VCA Shoreline
Veterinary Referral and Emergency Center
for evaluation as our emergency department
is open 24 hours a day. If emergency dialysis
is indicated based on the emergency doctor’s
evaluation, a member of the dialysis team
will be contacted. j
7
Clinical Laboratory Analysis:
Quality Samples Give Quality Results
Michele A. Papero, MHS, CVT
VCA Shoreline Veterinary Referral & Emergency Center
V
eterinarians today have many choices
regarding clinical laboratory testing
for their patients. The choice to use
a reference lab or perform tests in-house is
often based on a number of factors including
turn-around time, cost, equipment availability,
practicality and feasibility. Basic tests can often
be performed in the clinic, and until recently,
included only the most simple and cost-effective
tests. However, with the recent expanded
availability of in-clinic analyzers and rapid test
kits, many clinics are now performing a wide
array of diagnostic laboratory tests in-house.
Clearly, performing tests in the clinic has
advantages.
Results are available very
quickly and from fresh samples. However,
most clinics fail to maintain adequate quality
control and quality assurance programs
which seriously affects the integrity of
results. In addition, equipment and supplies
are costly to purchase and maintain often
making using a commercial laboratory the
less expensive option.
Large reference laboratories not only
provide an expansive test menu, but use the
most current technology, well-established
reference ranges, rigorous quality assurance
and quality control, employ highly trained,
credentialed personnel and offer consultation
services. However, there are also inherent
disadvantages to sending samples to an
outside laboratory. Turn-around time, while
usually within 24 hours for routine assays,
exceeds in-house testing times which may
delay diagnosis and treatment. Samples must
also be stored and transported for periods of
time which may affect their integrity.
protocols for acquiring, handling, preserving and
storing samples is critical as compromise in any
of these areas can seriously affect the integrity
of results and ultimately, jeopardize patient care.
Training of Personnel
It is imperative that all personnel asked to acquire
or analyze diagnostic samples be appropriately
trained. Technicians should understand the
difference between sample types, which tubes
are used for obtaining each sample type, how to
store samples, how to properly label samples, and
how to complete all paperwork and requisition
forms. Understanding the biology of a laboratory
test, even on a basic level, as well as the disease
state of the patient, will help technicians make
the correct choice when obtaining samples,
as well as identify results that do not seem
accurate. It is the responsibility of veterinarians
and senior staff to be sure that anyone asked to
obtain or process laboratory samples is trained
appropriately. Training should also include how
to find information so that sample processing
and analysis is not delayed, and how to maintain
and troubleshoot in-house analyzers. Many
test kits and reagents have specific storage and
handling requirements. Staff responsible for
running these assays should read and understand
all materials and instructions associated with the
assays and reagents as improper use and storage
will lead to compromised results.
Also important is understanding how to the
handle infectious and zoonotic samples. Every
clinic should establish protocols for obtaining,
storing and disposing of potentially infectious
samples in a way to prevent nosocomial transfer
between patients, and to protect employees.
Wound cultures suspected of containing MRSA
or other zoonotic bacteria, and urine samples
suspicious for leptospirosis are just two examples
of samples which may be infectious and when
improperly handled can lead to contamination
or illness. Other examples include blood and
cultures from patients suspected of having
Mycobacterium, nasal swabs or tracheal wash
samples from patients suspected of having
fungal infections, and scrapings from animals
with skin lesions that may contain mites, or
ringworm. The Centers for Disease Control
website (www.cdc.gov) has information sheets
on many zoonoses, which detail precautionary
measures that should be used when working
with samples in the clinic and laboratory. These
guidelines should be posted and followed by
any hospital staff member handling laboratory
samples for in-house or send-out analysis.
Packaging diagnostic samples for transport to
remote reference laboratories via courier service
Whether a hospital chooses to run samples
in-house or send out, the most important
considerations are the same. Sample acquisition,
sample handling and sample storage must be
flawless to guarantee sample quality and for
producing the most accurate results. When
obtaining samples, whether it is blood, urine or
tissue, it often helps to remember that the ideal
sample should reflect the in vivo state as closely
as possible. Establishing and maintaining strict
8
Full Circle Forum
such as FedEx or UPS and air transport should
be done to preserve the sample for the duration
of transport, and must be in accordance with
federal regulations. All clinical samples to
be transported by air courier service require
that special packaging and labeling be used to
comply with IATA (International Air Transport
Association) regulations. Both Federal Express
and United Parcel Service have extensive
information detailing the requirements for
transport of diagnostic clinical specimens on
their websites. Packages that are sent via FedEx
or UPS must be able to withstand jostling,
temperature extremes, and delays at transfer
stations or on delivery vehicles. Samples
should be packed so that the environmental
requirements recommended for the test to be
run are maintained. In some cases, special ice
packs and outer packaging may be required to
maintain the temperature inside the package.
Blood and other liquid samples should be in
containers that are secure and leak-proof, as tops
that are not secure can result in sample loss via
leakage and evaporation. Clinics can also receive
large monetary fines if packages show evidence
of leaking contents, or are not packed or labeled
correctly. It is the responsibility of the person
actually placing the sample inside the box to
be sure that all shipping regulations are met as
well as all of the requirements to ensure the safe
arrival of that sample at the laboratory.
Improperly labeled or packaged samples
sent using UPS or FedEx will be returned to
the shipper. In these cases, by the time the
time samples are returned, several days have
passed and the sample is no longer cold or
frozen and may have been exposed to extreme
temperature variations rendering it useless
and delaying important diagnostic results.
Assuring that all personnel are trained is the
best way to avoid this debacle.
Preparation for sampling
Personnel responsible for obtaining samples
for laboratory analysis must ensure that all
sample requirements are met including patient
preparation, timing of sample acquisition
(e.g. fasting, or timing after administration of
medication), and special handling (e.g. freezing,
centrifugation). One of the most common
errors is from submitting the wrong sample type.
An example of this would be submitting serum
for a test that requires plasma. Submitting one
type when another is required will render results
useless. Many drug assays have special sample
requirements, for example they may require
that blood or urine samples are taken during
a small window of time before the next dose of
medication or after the last dose of medication
March/April 2011
has been administered, or samples may require a
special tube. Serum for digoxin levels, for example,
should not be drawn into serum separator tubes
as the gel can interfere with recovery of the drug.
Taurine levels require plasma, not serum, and
must be kept cold at all times.
Periodic staff training sessions can be helpful
to make sure that everyone is following
the same protocols, and to review any new
information when it becomes available. There
are also times when a lab or assay kit adopts
new instructions; these should be reviewed
with staff and updated in a reference notebook
so that everyone is using the same set of
instructions. Most reference laboratories and
equipment manufacturers offer lunch and
learn sessions that can help keep the clinic
abreast of these changes and review hospital
laboratory protocols with staff.
While there are a variety of sample types
obtained in most clinics, blood and urine are
the most common. The following sections
make recommendations for acquiring good
quality samples for analysis.
Blood samples
Blood is, by far, the most common sample used
for diagnostic testing. Blood collection and
analysis provide a minimally-invasive means to
assess many physiological parameters. Relative
numbers of red blood cells, white blood cells,
and platelets are determined on the complete
blood count using anticoagulated whole
blood and these results are very important in
determining a patient’s well-being. As well,
serum analysis provides information on lipids,
enzymes, hormones, and antibodies. Whole
blood and serum sample integrity is dependent
on many things, and a poorly acquired or
handled sample has great potential to yield
skewed results on many tests. Below is a list
of considerations to ensure highest quality
samples are obtained for analysis.
1. Order of draw—To avoid cross
contamination of tube additives, collection
tubes should always be drawn in specific
order. For example, a serum sample will
show a very high potassium level when
there is contamination of the sample
with the anticoagulant potassium EDTA
contained in the purple top tube, which
can be transferred via the needle to the
serum separator tube. To avoid these
issues, tubes should be drawn in the
following order: Serum separator (gel
separator) or red top (no additive), blue
top (sodium citrate for coagulation assays),
green top (heparin), purple top (EDTA).
All sample tubes should be gently inverted
several times to mix the blood sample
with the anticoagulants or clot activators.
Never shake samples vigorously.
2. Phlebotomy—Personnel should be
experienced and proficient in drawing
blood. Most common lab tests require
venous blood however on occasion
arterial blood samples may be required
for tests such as blood gas analysis.
Special procedures and handling are
required for arterial blood analysis.
Arterial blood sampling requires
additional skill and careful attention
to avoid hemorrhage and hematoma
formation. Special collection syringes
and tubes are necessary so that blood
gases to be analyzed are not volatilized
prior to analysis. Arterial samples should
be analyzed immediately or stored cold in
an anaerobic environment to avoid errors
in blood gas analysis.
When obtaining venous blood samples,
superficial veins which are easily palpated
should be used. The animal should be kept
in a position where it can remain calm and
comfortable.
Excessive paw “pumping”
should be avoided as it can lead to hemolysis
which can alter serum chemistry results and is
undesirable for most tests. Avoid “probing” the
vein repeatedly as this can lead to hematoma
formation, and as a result, samples obtained
may contain unwanted clots. Prolonged
tourniquet application causes increased
pressure in the vein which leads to artifactual
changes in total protein, lipids, cholesterol, and
AST results. Drawing the plunger of a syringe
back too quickly when obtaining blood can
damage cells and also cause hemolysis.
Whenever possible, a 22g or larger needle
should be used for obtaining blood samples
as smaller gauge needles can lead to blood cell
damage. When transferring blood from the
syringe to the tubes, avoid forcing the blood
through the needle. Vacutainer needles and
butterflies allow the vacuum to draw the blood
into the tube. They are designed so that the
proper amount of blood is drawn into the tube
with minimal trauma to the cells. Never draw
blood samples from intravenous lines unless you
can withdraw and discard at least three times the
volume of the line before obtaining the blood
sample. Constituents of intravenous fluids and
medications administered through the IV line
can cause sample dilution and alter test results.
3. Sample quantity—There will always be
times when it is difficult to obtain sufficient
Continued on page 10
9
Quality Samples Give Quality Results
sample amounts (blood, urine, etc.) for
testing. Patient status (hydration, anemia),
demeanor, and many other factors can
preclude obtaining appropriate sample
quantity. However, both overfilling and
underfilling blood tubes can seriously alter
test results. Underfilling tubes containing
anticoagulants can lead to cell distortion,
altered red blood cell indices (MCV,
MCHC), falsely lowered hematocrit (in the
case of liquid EDTA) and falsely elevated
protein levels by refractometer. Citrate (blue
top) tubes containing insufficient sample
quantity leads to dilution of coagulation
factors and falsely prolonged clotting times.
At the same time, overfilling tubes will
overwhelm the anticoagulant and lead to
unwanted clotting of samples. All sample
tubes have the capacity in milliliters written
on the label. It is a good idea to order several
sizes of each tube type so that smaller tubes
can be used for smaller samples, providing
the final sample quantity is sufficient for the
tests requested. Outside laboratories, such as
Antech Diagnostics, provide a test manual
which lists the minimum quantity of sample
that can be submitted for each test.
4. Blood sample handling and storage—
Prior to obtaining blood samples, any
unique handling requirements should
be identified. Blood drawn into purple
top EDTA tubes should be refrigerated
immediately and can be submitted for up
to 72 hours if stored properly. Citrated
tubes for coagulation analysis should be
analyzed or refrigerated and submitted
as soon as possible. For serum separator
tubes blood should be allowed to clot,
with the tube upright, for 30 minutes and
then centrifuged immediately. Delay in
centrifugation will allow the blood cells
to continue to metabolize causing falsely
lowered glucose levels and alteration of
some hormone levels. Separated serum
for chemistry analysis can be stored in
Continued from page 9
Some clinics will accept samples in recycled
food containers or jars, however residual
detergents or food materials can affect the
results by adding glucose or altering pH.
Using opaque rather than clear containers
will protect the sample from direct
light which can lead to photochemical
breakdown of
clinically important
components such as bilirubin. When
possible, it is best to use commercially
available urine collection containers. These
can be disposed of, or cleaned, rinsed well,
dried and re-sterilized in ethylene oxide
gas. Patient information should never be
written on the lid of the container; always
label the container itself as lids can be lost
when the container is opened.
the refrigerator for 3-4 days. For assays
other than chemistries using serum, it is
best to check with the laboratory or the
manual for the analyzer if running inhouse. Some tests require that samples
be frozen immediately, however, repeated
freeze-thaw cycles can damage some
constituents. In this case, a common, selfdefrosting household freezer should not
be used. Occasionally, a test may require
protection from light or some other
special storage or handling procedure.
These requirements are usually listed in
the reference laboratory manual.
Urine samples
Analysis of urine is valuable in diagnosing,
monitoring, and screening for a number of
conditions and disease states. Some urine tests
require simply a “clean catch” of urine, and
others require samples be obtained sterilely
via catheter or cystocentesis. In either case,
there are many things that can affect results.
1. Containers—Containers used for
collection and transport of urine specimens
should be rigid, opaque and have a screw
top to minimize leakage and evaporation.
For samples which have been obtained by
cystocentesis into a syringe, never send the
syringe and/or needle when submitting.
The urine should be transferred, using
sterile technique, into a sterile red top tube
with no additives.
2. Sample handling and storage—Ideally,
urine should be analyzed within two hours
of collection, however, when sending to a
reference lab this is usually not feasible.
Therefore, to maintain sample integrity,
it is best to refrigerate urine samples
immediately to prevent proliferation
of bacteria which can alter glucose and
ketone concentrations. In addition, urine
stored at room temperature loses carbon
dioxide which will elevate the pH leading
to the formation or dissipation of crystals
over time. Many crystals are actually
soluble in alkaline urine, so those crystals
which could indicate pathology may not
be observed if the sample is not stored
properly. Refrigerated samples can be
kept and analyzed for up to 72 hours
according to our consultants at Antech
Diagnostics.
There are some commercially available
Pet Loss Support Meetings
First and third Thursdays of every month, Laurie Sine,
LMSW, 7 pm—8:30 pm at Animal Specialty Center, 9 Odell
Plaza, Yonkers, NY. Free of charge, open to the public.
Every Tuesday, Irene Javors, 7:30 pm at Fifth Avenue
Veterinary Specialists, One West 15th Street, New York, NY.
Free of charge, open to the public.
10
Full Circle Forum
Failure to perform this simple procedure can
result in seriously erroneous readings which
could affect diagnosis of disease state and
selection of treatment plans.
preservatives for urine, although they are not
recommended because while they preserve
cellular components, they alter the chemical
components of the sample. Some special
urine tests (e.g., catecholamines) may require
that the urine sample be quickly frozen for
preservation. Again, it is important to have
all of the information prior to obtaining the
sample so that results are not affected by poor
sample handling or storage.
Careful attention should be paid to
centrifugation speed when processing urines
for in-house analysis. Speeds that exceed
1500-2000 rpm can damage cells and crystals.
Refractometers used for measuring urine
specific gravity and total plasma/serum protein
should be calibrated weekly by placing a drop
of distilled water on the glass and zeroing the
device following the manufacturer’s directions.
VCA Shoreline and VCA VREC are pleased to announce the addition of
VCA SPECIALTY IMAGING SERVICES
Our specialists are now available to travel to your office in Southwestern Connecticut.
Services include echocardiography, ultrasound imaging, consultation, case review and
discussion of therapeutic recommendations with you about each case.
Echocardiograms
Nathaniel Fenollosa, DVM, DACVIM (cardiology)
Abdominal & Thoracic Ultrasounds
Larry Berkwitt, DVM, DACVIM (internal medicine)
Dianne Kittrell, DVM, DACVIM (internal medicine)
Beth Whitney, DVM, DACVIM (internal medicine)
Lisa Keno, DVM, DACVIM (internal medicine)
Michelle Cieplucha, DVM (practice limited to internal medicine)
Using our GE Vivid i portable ultrasound system, we will be able to perform inoffice echocardiograms, abdominal, and thoracic ultrasounds.
The Vivid i is a portable cardiovascular ultrasound unit which allows us to obtain
real-time diagnostic information. Our new system is expanded not only for high
performance echocardiography, but is optimized for general, high-resolution
imaging of the thoracic and abdominal cavities, and vasculature.
If you are interested in scheduling any of the above specialists to visit your office, or
to obtain more information about this service, please contact Dr. Larry Berkwitt or
Michele Papero at (203) 929-8600, or Dr. Sam Durkan or Kay Wyler at (203) 854-9960.
We very much look forward to working with you on this new venture!
March/April 2011
Dipsticks allow for analysis of the chemical
components of urine. Hemoglobin, pH,
bilirubin, ketones, glucose, protein, and
urobilinogen can all be semi-quantified on
a single dipstick. Each small pad on the
stick is impregnated with reagents. Chemical
constituents of urine are detected via a chemical
reaction and resulting color change on the
pad. These reactions are time dependent and
a common lab error is reading these color
changes too soon. Careful attention should
be paid to the recommended incubation times
prior to reading and recording the results of
each test on the strip.
While blood and urine are the most common
samples analyzed both in the clinic and reference
laboratory, other types of patient samples
include biopsy tissue, various body fluids, and
a variety of sample types used in molecular
biological testing like PCR. These samples
require very special handling or processing to
prevent degradation of DNA or damage to
cells and tissues. Improperly handled or stored
cultures will result in overgrowth or the loss of
bacteria and invalid results.
It would be impossible to address all of
these issues here especially since sample
requirements often depend on the method
of analysis used which may vary from lab to
lab. In these cases, contact your reference
laboratory directly to be sure samples are
obtained, handled, and submitted seamlessly
and to ensure the best opportunity for
accurate and timely test results.
Consultants at Antech Diagnostics are always
helpful, can answer all of your questions, and
can be reached at (800) 872-1001. j
References Used:
Fischbach F. A Manual of Laboratory and
Diagnostic Tests. 5th ed. Philadelphia:
Lippincott; 1996.
Rodak BF. Diagnostic Hematology.
Philadelphia: WB Saunders Company;
1995.
Willard MD, Tvedten H, Turnwald
G. Small Animal Clinical Diagnosis by
Laboratory Methods. Philadelphia: WB
Saunders Company; 1999.
Personal communication with consultants at
Antech Diagnostics.
11
Upcoming Events
March 24, 2011 — Animal Specialty Center CE
Program. Topic: Neurology: Updates on Seizures,
Jason Berg, DVM, DACVIM (Neurology/Internal
Medicine). Location: 9 Odell Plaza, Yonkers, NY
10701, 6:30 p.m. dinner, 7:30 p.m. lecture, RSVP:
(914) 457-4023 or [email protected].
March 29–30, 2011 — Please join VCA Shoreline,
VCA VREC, Animal Specialty Center, VCA Boston
Road Animal Hospital and VCA Cheshire Animal
Hospital at the upcoming Connecticut Veterinary
Medical Association Annual Meeting which has been
rescheduled for Tuesday, March 29 and Wednesday,
March 30 at the Mystic Marriott in Groton, CT. VCA
Specialists will be available for complimentary case
consultation and radiography review. Bring any
case materials and radiographs you would like to
discuss. For more information about the meeting
and how to register go to www.ctvet.org or email:
[email protected].
Full Circle Forum
One West 15th Street
New York, NY 10011
(212) 924-3311
(212) 924-7228 (fax)
April 28, 2011 — Animal Specialty Center CE
Program. Topic: Radiation Oncology: Update
on CyberKnife, Sarah Charney, DVM, DACVIM
(Oncology), DACVR (Radiology). Location: 9 Odell
Plaza, Yonkers, NY 10701, 6:30 p.m. dinner, 7:30
p.m. lecture, RSVP: (914) 457-4023 or rcc@
animalspecialtycenter.com.
May 17, 2011 — Fifth Avenue Veterinary Specialists will
host an evening of CE at 7:00 p.m. at the Union Square
Ballroom in NYC. Our featured speakers will be Dr. Kate
Margalit, DACVS, and Dr. Jessica Chavkin, DACVIM.
Please contact Monica Dunn at (212) 924-3311 or
[email protected] with any questions.
May 26, 2011 — Animal Specialty Center CE
Program. Topic: Surgery: Fluoroscopic assisted
minimally invasive fracture repair with Dennis Ting,
DVM, DACVS. Location: 9 Odell Plaza, Yonkers, NY
10701, 6:30 p.m. dinner, 7:30 p.m. lecture, RSVP:
(914) 457-4023 or [email protected].
ANNOUNCEMENTS
Dr. Matthew
Palmisano, staff
surgeon at VCA
Veterinary Referral
and Emergency Center
in Norwalk, CT has
been named Associate
Editor for Small Animal Orthopedics in
Veterinary Surgery. Veterinary Surgery
is the official publication of both the
American and European College of
Veterinary Surgeons. The magazine
is one of the foremost publications for
small and large animal surgery, and has a
worldwide distribution.
Fifth Avenue
Veterinary Specialists
is pleased to announce
that Dr. Kate Margalit
is now a Diplomate of
the American College
of Veterinary Surgeons.
We are proud to have her as part of the
FAVS specialty staff and look forward to
a long future of working with her.
First Class Prsrt.
US Postage
PAID
Canoga Park CA
Permit #451
Administrative Services provided
by VCA Animal Hospitals, Inc.