Following are differential diagnoses for patients presented with hyperglobulinemia. Hyperglobulinemia can be caused by monoclonal or polyclonal gammopathies; serum electrophoresis is required for differentiation and can help prioritize possible diagnoses. Polyclonal gammopathies are composed of nonalbumin proteins (ie, globulins) and are typically caused by inflammation, infection, or immune stimulation. Monoclonal gammopathies typically result from production of a single type of globulin protein and are most commonly associated with neoplastic causes, although rare non-neoplastic causes have also been described.

• Acute-phase reactant response (ie, tissue injury of any cause [eg, inflammation, acute bacterial or viral infection, necrosis, neoplasia, trauma]; typically mild)*
• Chronic antigenic stimulation/inflammation*
  • Bacterial endocarditis
  • Chronic skin disease
  • Immune-mediated disease (eg, systemic lupus erythematosus, immune-mediated hemolytic anemia)
  • Infectious disease (eg, FIP, leishmaniasis, heartworm disease, coccidioidomycosis, ehrlichiosis, hepatozoonosis, pythiosis, bartonellosis)
  • Liver disease (eg, lymphocytic cholangitis)
  • Severe dental disease
• Hemoconcentration (concurrent increase in albumin)
• Nephrotic syndrome*
• Paraproteinemia (due to abnormal immunoglobulin production resulting in a monoclonal gammopathy)
  • Infectious disease-associated monoclonal gammopathies (usually immunoglobulin G; eg, Dirofilaria immitis, Ehrlichia canis, visceral leishmaniasis)
  • Inflammatory disease (eg, lymphoplasmacytic enteritis, cutaneous amyloidosis; rare)
  • Neoplasia
    • Chronic lymphocytic leukemia
    • Extramedullary plasmacytomas that affects the skin (dogs), GI tract, or liver
    • Lymphoma
    • Multiple myeloma
    • Waldenström macroglobulinemia

SARS-CoV-2 is responsible for the global COVID-19 disease pandemic. The Coronaviridae virus family includes many important veterinary pathogens classified as alphacoronaviruses (eg, transmissible gastroenteritis virus, FIP, canine enteric virus) and betacoronaviruses (eg, SARS-CoV-2, bovine coronavirus, canine respiratory coronavirus). Companion animal coronaviruses are common pathogens and appear to be species-specific. Infection of respective hosts typically does not cause severe disease; most cases are mild or subclinical, except in the very young. These companion animal coronaviruses do not share antigenicity with SARS-CoV-2; thus, current vaccines for feline, canine, and bovine coronaviruses do not provide protection against SARS-CoV-2.

Feline coronavirus (FCoV) is also associated with FIP, which is a serious disease of cats that demonstrates a complex pathogenesis involving virus, host, and environmental factors. Although FCoV infection is not uncommon, FIP is rare and affects a minority of cats infected with FCoV. Neither FIP nor its causative virus (ie, FCoV) share pathogenic or antigenic properties with SARS-CoV-2.

SARS-CoV-2 is believed to have originated in bats and infected an intermediate host(s); this is similar to the association between civets and SARS-CoV-1. SARS-CoV-2 uses angiotensin-converting enzyme 2 (ACE2) to attach to and enter the cell. This molecule is potentially recognized by SARS-CoV-2 in a range of animal species (eg, palm civets, pigs, cats, ferrets, nonhuman primates), suggesting the virus may infect these animals and warranting further study.¹ It is therefore important that clinicians exercise precautions with any animal that has been in contact with an infected human.² See recommendations provided by WSAVA here.

It is also important for clinicians to reassure owners that the risk for contracting COVID-19 disease from their pet is currently determined to be low and that pets should not be euthanized, abandoned, or otherwise removed from households. Current recommendations for owners include²:

• Companion animals should be kept with owners who are self-quarantining. 
• Good hygiene practices, including regular hand washing, should be maintained when interacting with pets.
• Care for companion animals with family or friends should be arranged if owner hospitalization is required.
• A veterinarian should be contacted immediately with questions or concerns.

In addition, clinicians must protect their clinic and staff, including implementing strict protective practices, as with any respiratory disease patient. It is also important that disseminated information be valid and accurate. There is currently no evidence that domestic animals (eg, dogs, cats) can transmit COVID-19 to uninfected humans, and limited reports exist regarding clinical infection in dogs and cats.³   

The public will continue to look to the veterinary community for information and recommendations, and it is imperative that clinicians remain informed and serve as a resource for owners with concerns about pet-associated risks for infection. The veterinary community must provide reassurance and support for the welfare of patients and their owners. Objectivity and equanimity are of critical importance in times of stress.

This condition is commonly seen in large- and giant-breed dogs (eg, German shepherd dogs, Great Danes, mastiffs, Saint Bernards, Newfoundlands, Irish wolfhounds) that are 6 to 18 months of age.^4-7 Hygromas can occur over any bony prominence (eg, greater trochanter, ischiatic tuberosity) but most commonly occur over the olecranon of the elbow.^2,3,8

Elbow hygromas can be classified as uncomplicated or complicated based on clinical appearance.⁹ Uncomplicated elbow hygromas are small, painless, and nonulcerated and can progress in severity as trauma occurs and continues, resulting in cellular death, ischemia, and edema. Complicated elbow hygromas are typically recurrent, large, painful, ulcerated, and/or infected.⁶ Calcinosis cutis circumscripta-type lesions appearing secondary to chronic hygromas have also been described.¹⁰ Other classification schemes for elbow hygromas based on the mechanisms of development have been reported.⁵

Conservative management of uncomplicated elbow hygromas aims for resolution in 2 to 3 weeks; fibrous connective tissue can develop during this healing process.^3,4 Aspiration of the hygroma fluid may be attempted using an aseptic technique, but recurrence and infection are common sequelae.^3,4 Intralesional corticosteroid administration is not recommended because of the risk for acute infection following injection.⁸ Conservative management includes educating pet owners about at-risk breeds and emphasizing the importance of maintaining the patient’s ideal body weight, protecting elbows from hard surfaces using soft bedding, and maintaining loose padding of the elbows, especially in early stage of elbow hygroma development.^1,4 Although neoprene/polyester-padded elbow sleeves are commercially available, their clinical efficacy has not been evaluated.

Surgical correction should be reserved for complicated elbow hygromas and can include:

• Passive or active drainage of cavitated lesions with external coaptation⁶
• Releasing incisions or using tension-relieving closing techniques^2,3,11
• Ostectomy of the olecranon to decrease pressure on the overlying skin³
• Reconstruction after excision of the hygroma using single pedicle advancement flaps, single pedicle direct flaps, and axial pattern flaps (eg, superficial brachial artery, thoracodorsal artery, or rectus abdominis free muscle flaps)^3,12
• The microvascular free muscle transfer technique⁷
• Surgical repair with commercially available foam pipe insulation for the protection of elbows¹³
• Surgical debridement and vacuum-assisted closure with application of negative pressure wound therapy (NPWT)¹⁴

Postoperative external coaptation with soft padded bandaging or splinting is commonly used to immobilize the elbow joint, regardless of surgical technique. NPWT has been used in wound management in small animal patients and can be useful for large unhealthy wound beds.¹⁵ NPWT aids in closing wounds through development of healthy granulation tissue, which enables later primary wound closure. NPWT has been shown to improve the wound healing process, decrease the frequency of bandage changes, and decrease the risk for contamination. Limitations include equipment, labor costs, and the need to train staff members, including clinicians.

Complications reported with surgical correction of elbow hygromas include skin dehiscence associated with tension, surgical site infection, splint- and bandage-related lesions, seroma formation, pain, delayed healing, and, with NPWT, loss of the periwound seal.^4,8,14

Orthogonal elbow radiography is recommended prior to surgery to rule out osteomyelitis, periostitis, and neoplasia as possible causes of hygroma formation.³ Sedation may be required for tissue biopsy and deep cultures prior to surgery if the hygroma is complicated and ulcerated (Figure 1).

Postoperatively, the patient should be hospitalized for 3 to 4 days and monitored at least every 2 hours for fluid production and to ensure consistent negative pressure. Alternatively, the patient can be discharged with the unit for at-home monitoring (Figure 2). Owners should be educated about the unit so they can troubleshoot if necessary and be provided with multiple canisters so they can change the canister as it fills.

NPWT can be discontinued 3 to 4 days postoperatively with the patient under general anesthesia (Figure 3). The resultant wound bed may have evidence of stimulated and evenly distributed granulation tissues, allowing for delayed primary closure with interrupted and tension-relieving (near-far-far-near pattern) skin sutures (3-0 nylon).

External coaptation should be performed for an additional 10 to 14 days until the incision heals (Figure 4). Bandages should be changed every 3 to 5 days to assess the wound for complications. As an alternative to external coaptation, foam insulation can be applied to protect the elbow.¹³ Once the surgical site heals, the owner should be advised of the necessary lifelong changes to care, including providing soft bedding, padding the elbows if erythema is observed (usually the first sign of pressure sores), and maintaining an ideal body condition.

• Standard surgical pack with #10 or #15 scalpel blades
• Sterile isotonic saline
• Monopolar electrocautery or Mayo or Metzenbaum scissors
• Polyurethane ether foam dressing and vacuum pad
• Vacuum-assisted closure system, including canisters with attached tubing
• Semipermeable adhesive drapes
• Antimicrobial incise drapes or transparent film dressing
• Medical adhesive spray or stoma paste
• Nylon suture (3-0)
• Bandaging material and tape

Clip the affected thoracic limb from mid-dorsum of the cervical region distally to the digits. Position the patient in lateral recumbency with the affected limb isolated and the lateral side exposed. Aseptically prepare the limb while it is hung from the ceiling or other device. Perform standard draping.

Administer perioperative IV antimicrobials (eg, cefazolin) 30 minutes prior to making an incision; re-administer every 90 minutes. Make a proximal-to-distal incision using a #10 or #15 scalpel blade over the wound, incorporating the open ulcerated wounds. Carefully dissect the SC and/or necrotic tissue directly and sharply with monopolar electrocautery or Mayo or Metzenbaum scissors. Simultaneously perform surgical debridement of any necrotic tissue.

Copiously lavage the wound site with warm sterile isotonic saline prior to closure. Obtain a tissue sample for bacterial culture and susceptibility testing if needed.

Apply foam dressing to the wound bed; avoid overlapping skin to prevent skin maceration and damage. Apply adhesive spray or stoma paste 3 to 5 cm around the periwound skin as needed. Next, apply a semipermeable adhesive drape to the entire wound bed, covering the skin and foam. Using a #10 scalpel blade, make a 2-cm slit through the drape over the foam, remove the protective covering from the vacuum pad, and attach the vacuum pad to the slit area. Direct the tubing toward the dorsum, not distally. Cover the pad with another layer of semipermeable adhesive draping.

Attach the suction tubing to the collection canister and initiate the pump unit. Maintain pressure at -125 mm Hg.¹³ After the pump is activated, confirm successful dressing placement through visualization of shriveling, hardening, and wrinkling of the foam and drapes.¹⁴

Facial trauma from motor vehicles, bites, or falls can result in maxillary and/or mandibular fractures. These fractures represent 2% of all fractures in dogs and 15% of those in cats.^1,2 In older patients, bone resorption can lead to self-induced or iatrogenic fractures with minimal trauma during dental procedures.

Patients with jaw fractures have typically experienced recent motor vehicular trauma, animal bites, blunt force trauma, or falls (including high-rise syndrome). Some fractures may not be evident to the pet owner until food prehension difficulties arise. Many fractures are discovered based on the patient’s inability to close the mouth, visible malocclusion, pytalism, and/or bleeding from the mouth.

A thorough oral examination with the patient under general anesthesia is the most straightforward diagnostic technique. Palpation of the symphysis, horizontal body of the mandible, vertical ramus, temporomandibular joint, maxillary arcade, and hard palate is the first step to determine further diagnostics. Whereas open fractures can usually be observed, more caudally located fractures can be difficult to observe; presence of blood in the mouth should raise suspicion. Symphyseal separation is typically straightforward to palpate; however, the clinician should always look for a second fracture using general or dental radiography. Standard orthogonal views may be supplemented with oblique views to reduce confusion due to superimposition of teeth and bones. The maxilla can be assessed with these methods but is often best evaluated with CT if multiple fracture areas are suspected. Radiography can be used to evaluate fracture orientation, fracture number, tooth viability, bone quality, osteomyelitis, and tumor formation.

Other body systems should be assessed for damage secondary to any primary trauma. Common injuries include brain trauma, cranial nerve damage, and pneumothorax; thus, a thorough physical examination and cardiopulmonary stabilization should be performed prior to treatment for facial fractures.

General anesthesia is required for fracture treatment, and local anesthetic blocks are advantageous. IV antibiotics should be administered in patients that have open fractures. Intubation with a short endotracheal tube and pharyngostomy are useful for evaluating occlusion during fracture surgery; in the former, the connection between the tube and anesthesia hose is located in the mouth, allowing for brief, intermittent disconnection with closure of the mouth to assess interdigitation of the maxillary and mandibular teeth. A pharyngostomy tube can be placed caudal to the mandible through a separate skin and pharyngeal incision (Figure 1) or through an incision ventrolateral to the tongue.³

Symphyseal separation can be repaired by passing 22- to 18-gauge wire through a hole in the ventral midline skin to encircle the base of the mandibular canine teeth (Figure 2). The wire should be tightened to ensure stability of the symphysis while occlusion is observed to maintain proper spacing between the mandibular and maxillary canine teeth. The cerclage wire should be cut with sufficient wire protruding to enable removal after 6 to 8 weeks. The metal twist may be covered by a dollop of bone cement, acrylic, or a pencil eraser to prevent self-trauma.

When managing fractures of the mandibular body, the surgeon’s goal is to provide stability to achieve bone union, whereas the dentist’s goal is to maintain tooth viability and proper occlusion. For example, a surgeon may want to place cerclage wires near the tooth root in the ventral mandible to secure good-quality bone at the tension surface, whereas a dentist might prefer an intraoral technique to preserve the root and relinquish a biomechanically superior location for implant placement. With proper planning, these goals can be balanced with a variety of methods. Interfragmentary wiring can be performed by placing 24- to 18-gauge wire throughout the length of the mandibular body; typically, >1 wire should be placed perpendicular to the fracture line for stability (Figure 3). Simple straight-line fracture configurations are best for this technique. In cases involving comminution of the mandible, plates or external fixators can be useful (Figures 4 and 5). These are typically placed on the lateral aspect of the mandible with careful avoidance of the tooth roots.

When fracture lines between intact teeth are observed, interdental wiring with 26- to 20-gauge wire at the gingival line of each tooth can be useful as a supplement to other techniques. Acrylic is commonly added over the teeth–wire construct. The teeth should first be scaled, polished, and acid etched, then acrylic should be applied over the teeth–wire construct while ensuring occlusion (Figure 6).^4,5 Bone healing may be prolonged if tooth extraction or root canal is required.⁴ The acrylic and wires can be removed once bone healing is evident, typically 6 to 8 weeks after injury.

In cats and small dogs with caudal body or ventral ramus fractures, bonding of the 4 canine teeth is a viable treatment option. Bonding prevents the entire mandible from moving so that a fracture in any location will heal. It is most often used with caudally located fractures for which there are fewer repair options. However, the application often gets overpowered and fails in medium and large dogs, for which other methods are needed. The canines should be scaled, polished, and acid etched, then acrylic should be applied while the mouth is open in a manner that would enable intake of gruel. The acrylic should be removed in 6 to 8 weeks. In older dogs with small rostral mandibular fractures, partial mandibulectomy is an excellent treatment strategy with no potential for nonunion or bone infection.

When financial considerations preclude the fracture assessment and treatment plan, a tape muzzle can be used. The face should be cleaned and dried and any open fractures sutured closed. Tape should then be applied in a circular pattern over the muzzle to allow an oral opening sufficient for intake of gruel. Another piece of tape should be placed under the ears and around the back of the head and secured to the muzzle tape (Figure 7). An Elizabethan collar should be used to prevent the patient from removing the muzzle. Pet owners should be advised to clean the tape muzzle, although it can be easily replaced if necessary. A sufficiently large nylon muzzle may alternatively be used.

Postoperative treatment includes pain management (eg, fentanyl or hydromorphone immediately, followed by an oral NSAID with tramadol or gabapentin) and 1 to 2 weeks of antibiotic treatment (eg, amoxicillin/clavulanic acid, clindamycin) for open fractures. In addition, the patient should be provided soft food for 2 to 4 weeks, during which time toys and hard treats should be withheld. Follow-up radiography should be performed 6 to 8 weeks posttreatment to assess fracture healing. Jaw fracture treatment has a complication rate of 34% to 60%.^1,2 Malocclusion and infection are the most common complications; implant failure, malunion, and nonunion may also occur. Nonunion can be treated with further stabilization surgery and bone grafting.

Evaluation of pet owner purchasing patterns of flea and tick prevention has shown that owners rarely purchase enough medication to provide consistent, year-round coverage for their pet.²

In a recent study, >500 dog owners in 24 clinics across the United States were surveyed²; despite recommendations for year-round prevention, only 73% of owners believed that year-round prevention was necessary. The average owner bought only 6.1 months of prevention, with only 13% of owners purchasing a year’s supply of medication.

Even after purchasing medications with the best of intentions, owners may fail to administer preventives, which may occur for many reasons (eg, forgetfulness, confusion regarding flea and tick seasonality, difficulty administering medication).

Educating owners on the importance of regular, consistent flea and tick prevention is an essential step to achieving compliance and overcoming purchasing barriers.¹ Proper education, including messaging from the entire practice team, can be built into routine appointments to reinforce the importance of flea and tick prevention. Perception of necessity, along with cost, has been shown to be vital to follow-through in human medication puchases.³

Successful education efforts should be followed up by sending owners home with preventives after every routine appointment. Convenience can address key barriers to administration. Studies have shown that inconvenient routes of administration and more frequent administration are associated with poor compliance, whereas less frequent administration has been associated with higher compliance rates.^4-6 Preventive products that have convenient dosing and administration may promote successful and timely administration, improving compliance once the medication is purchased.³ Seresto®, a slow-release imidacloprid/ flumethrin collar that provides flea and tick prevention for 8 months, can help break the barriers of cost, convenience, and ease of administration, all while maintaining high standards of efficacy.^7,8 Collar placement is simple and can be done during the appointment, eliminating any potential owner concerns regarding proper placement. Mid-year rechecks can also be encouraged to alleviate owner concerns regarding potentially missing prevention doses and help owners feel confident that prevention is under the supervision of the clinician. 

Clinicians must recognize that compliance barriers may cause flea and tick prevention to fall short of their expectations and that these barriers will vary among pet owners. Clinicians should communicate with owners to identify the individual barriers and tailor their recommendations to overcome them. By employing concerted education efforts and preventive products that remove the guesswork of compliance, clinicians may turn their compliance expectations into realities. 

On examination, Trooper was bright, alert, and responsive. All peripheral lymph nodes were normal. Periocular regions were alopecic with mild to moderate crusting (Figure). The pinnae were moderately erythematous on the concave surface with mild adherent scaling. The scrotum was severely erythematous. All paw pads were crusted. The remainder of the examination was unremarkable.

Evaluation of an impression smear of the periocular region revealed numerous cocci with streaming neutrophils. A deep skin scrape was negative for ectoparasites. Skin biopsies were discussed with Trooper’s owner; however, superficial pyoderma was initially treated prior to biopsy to ensure accurate results. Bacterial culture and susceptibility testing was performed on the lesions to guide therapy.

Four days after initial presentation, culture results revealed methicillin-resistant Staphylococcus schleiferi (Table). Treatment with chloramphenicol (40 mg/kg every 8 hours) was initiated. In addition, the owner was instructed to bathe Trooper weekly using a 3% chlorhexidine/phytosphingosine shampoo. Due to the location of the periocular lesions, topical therapy did not seem appropriate as the sole means of eliminating pyoderma.

Infection had improved significantly 2 weeks after initial presentation, and Trooper was returned to the clinic for punch biopsies (6 mm) of the affected areas, performed while he was under sedation. The specimens were submitted for histopathology, which confirmed zinc-responsive dermatosis (see Histopathology Results of Punch Biopsies Indicating Marked Parakeratotic Hyperkeratosis).

Chloramphenicol therapy was continued for 4 weeks, including 1 week after resolution of clinical signs of infection and cytologic cure. Supplementation with zinc methionine (elemental zinc, 3 mg/kg once daily [dose can be split and given every 12 hours]) was initiated (see Treatment at a Glance).^1-3

Trooper was rechecked 1 month after zinc therapy was initiated; lesions showed some improvement, and evaluation of an impression smear confirmed the absence of bacteria. Low-dose methylprednisolone (initial dose, 0.7 mg/kg once daily) was initiated and tapered over 1 month. In many cases, low-dose corticosteroids can be beneficial for treatment of dogs that do not respond to zinc alone,⁴ as corticosteroids are known to increase zinc absorption from the GI tract.

Trooper was presented 6 weeks later for a recheck examination. He had not received methylprednisolone for 2 weeks, and all lesions were resolved. Zinc methionine supplementation was continued, and Trooper was free of lesions. Lifelong zinc supplementation is typically needed. Long-term prognosis is favorable as long as zinc supplementation is maintained.

Zinc-responsive dermatosis syndrome I is a condition that occurs primarily in Alaskan malamutes, Siberian huskies, and other arctic breeds and may be associated with defective intestinal absorption of zinc.^1,2,5

Syndrome II occurs in dogs fed a zinc-deficient diet.^1,2,6 The author has anecdotally seen an increase in the number of syndrome II cases that may be a result of an increase in dogs being fed home-prepared and alternative diets. Lesions are typically located at mucocutaneous junctions and paw pads and appear as areas of erythema with scaling, crusts, and hyperkeratosis.

Zinc-responsive dermatosis should be on the differential diagnosis list for crusted skin disease in any northern- or arctic-breed dog (see Take-Home Messages). Diagnosis is made through history and biopsy. Serum or hair levels of zinc may be low in affected patients; however, proper analysis can be difficult due to a variety of factors, so biopsy is the recommended diagnostic test if zinc-responsive dermatosis is suspected.⁷

Osteoarthritis (OA) and chronic kidney disease (CKD) are common in older cats.¹ It has been suggested that OA-associated pain and reduced mobility cause decreased water consumption, leading to worsening prerenal azotemia, constipation, and, ultimately, CKD progression. NSAIDs can decrease lameness in cats with OA; however, use of NSAIDs to manage OA pain in cats with CKD has historically been discouraged.² NSAID administration reduces the renal production of prostaglandins, which are important regulators of glomerular pressure, sodium reabsorption, and renal perfusion. Blocking their production may precipitate renal injury, particularly if a cat is hypovolemic or dehydrated.³

This article review by the WSAVA Global Pain Council evaluated recent studies that examined whether meloxicam and robenacoxib can be safely administered to cats with CKD. The 3 clinical studies referenced had generally favorable outcomes, although limited study duration, reduced drug dosages, and case selection biases limit the broad application of the results. In the studies, adverse event frequency and lifespan were similar between the meloxicam/robenacoxib-treated and control groups.^4-6 In one study, cats receiving meloxicam experienced a slower increase in median serum creatinine over time⁴; it is unclear whether this effect was due to increased mobility and reduced pain, allowing for increased water and food consumption, or possibly due to reduced tubulointerstitial inflammation. These study conclusions are also supported by a cat remnant kidney model study in which euvolemic cats with experimentally induced azotemia did not experience changes in glomerular filtration rate after short-term meloxicam administration.⁷

Despite these findings, NSAID administration in cats with CKD requires careful consideration based on patient stability and owner education. In cats, NSAIDs can cause acute kidney injury, and, in hypovolemic dogs, their administration decreases renal function.^3,8 To mitigate these risks, the WSAVA Global Pain Council recommends NSAID doses be tapered to the minimal effective dose. In addition, NSAIDs should be avoided in cats with progressive azotemia or weight loss and used cautiously, if at all, in cats with International Renal Interest Society stage 3 or 4 CKD, considering the lack of clinical data in this patient population. Before and during therapy, comorbidities (eg, dehydration, proteinuria, hypertension, hyperphosphatemia) must be carefully managed and regularly monitored. At home, owners must continually assess their cat’s water and food intake, changes in weight, and clinical signs. Optimizing care with alternative therapies (eg, environmental enrichment, physical therapy, acupuncture, and nutraceuticals, including chondroprotective agents) is recommended.

Key pearls to put into practice:

Meloxicam or robenacoxib can effectively manage OA pain in cats that have stable, well-managed stage 1 or 2 CKD.

Titration to the lowest effective NSAID dose is recommended in combination with other nonpharmaceutical therapies for OA pain.

Meloxicam or robenacoxib administration carries risk. Owners should be educated about these risks and be active participants in monitoring their cat. They should understand that dehydration from decreased food/water intake or increased losses (eg, vomiting, diarrhea, polyuria) in combination with NSAID use is dangerous and can cause acute kidney injury with a rapid decline in kidney function.

The pelvis is the second most common fracture site in cats, with ≈25% of all reported fractures occurring in the pelvis.^1-3 The most common causes include falls from a tall height and vehicular trauma.² Concurrent extra-pelvic injuries are also reported in 58% to 72% of cats with traumatic pelvic fractures, as large impact forces tend to be involved with these injuries.^2,4 

In humans with traumatic pelvic fractures, the mortality rate is 5% to 50%,⁵ depending on overall severity of injury; ≈33% of patients with traumatic pelvic fractures require blood transfusion as part of resuscitation.⁶ Most occurrences of hemorrhage in humans are believed to stem from concurrent injuries, not from fractured bone.^5-8

The need for transfusion and hemorrhage-control interventions in humans can be predicted by the pattern of pelvic fractures, the presence of shock on admission, and the Injury Severity Score.^9,10 This retrospective study evaluated whether characterization of transfusion requirements and outcomes could be similarly predicted in cats presented with traumatic pelvic fractures. Of the 112 cats included in the study, 21 received a blood transfusion, 84 required surgical fracture stabilization, 25 required surgery for other injuries (ie, skin wounds, urinary tract trauma, ocular trauma), and 102 (91.1%) survived to discharge. Only half of the cats requiring a transfusion needed it preoperatively, and none received the transfusion as part of initial resuscitation.

Cats with sacroiliac luxation or pubic fractures were more likely to receive blood transfusions; however, these fractures were also the most common, so further evaluation in a different population of cats is needed to determine if these fracture types can truly predict the need for transfusion. Of note, 8% of cats in this study required surgery to repair disruption of the urinary tract, a rate much higher than previously reported in either dogs or cats.^2,7,8

Key pearls to put into practice:

Approximately 1 in 5 cats with pelvic fractures may require a blood transfusion during hospitalization.

Because cats with traumatic pelvic fractures often have additional injuries, any cat with a pelvic fracture should have a thorough, whole-body-systems evaluation.

Urinary tract injury may be more common in cats with traumatic pelvic fracture than previously thought. Contrast cystourethrogram and/or serial ultrasonography may be helpful for early identification of cats requiring intervention for this problem.

In cases of severe hypoxemia, use of a traditional nasal cannula or oxygen-enriched environment may not be sufficient to support oxygenation, in which case mechanical ventilation may be necessary.¹ However, given the need for specialized equipment and expertise, mechanical ventilation may not be practical in many clinical situations. The opportunity to provide higher levels of supplemental oxygen and continuous positive airway pressure (CPAP) may allow for respiratory support without mechanical ventilation.

High-flow nasal cannula (HFNC) oxygen delivery systems have been developed and used in human medicine with the goal of delivering much higher flows of oxygen, which are better tolerated than traditional flows as the systems heat and humidify the air. In addition, the tight seal and high flow of oxygen allow the generation of CPAP, which mimics the effects of positive end expiratory pressure achievable with mechanical ventilation. Both CPAP and positive end expiratory pressure can help improve pulmonary function by decreasing atelectasis and promoting lung recruitment. This research study sought to determine the safety and efficacy associated with application of an HFNC system to dogs.

A total of 8 healthy dogs were included in this randomized crossover study. Study groups included traditional nasal cannula (at 0.1, 0.2, and 0.4 L/kg/min flow rates) and HFNC with subjects either awake or sedated (at 0.4, 1, 2, and 2.5 L/kg/min flow rates). Measured parameters included inspiratory and expiratory airway pressure, fraction of inspired oxygen (FiO₂), partial pressure of oxygen, partial pressure of carbon dioxide, temperature, heart and respiratory rate, arterial blood pressure, and pulse oximetry. Complications and predefined tolerance and respiratory scores were also assessed.

The HFNC junior interface fit well on 3 dogs; however, the adult interface had to be modified to fit well on the other 5 dogs. No differences were found with regard to vital parameters between the traditional nasal cannula and HFNC groups. The HFNC group showed good tolerance at 0.4 and 1 L/kg/min, acceptable tolerance at 2 L/kg/min, and poor tolerance at 2.5 L/kg/min, with CPAP being achieved at flows ≥1 L/kg/min. Dogs in the traditional nasal cannula group receiving 0.1 L/kg/min failed to have an increase in FiO₂ but achieved an average of 50% at 0.2 L/kg/min and 72% at 0.4 L/kg/min. With HFNC, FiO₂ averaged 72% at 0.4 L/kg/min and 95% for all other flow rates assessed, with minimal impact on ventilation. Dogs receiving HFNC showed radiographic evidence of aerophagia, but no other complications were noted.

Key pearls to put into practice:

Although previously reported to be effective,¹ traditional nasal oxygen supplementation with 0.1 L/kg/min failed to achieve FiO₂ statistically different from room air in this study. Target levels of 0.2 to 0.4 L/kg/min should be considered.

HFNC oxygen therapy is well-tolerated at rates of 0.4 L/kg/min to 2 L/kg/min and can achieve CPAP at flows ≥1 L/kg/min with no significant complications. Nasal cannulas may need to be modified for medium- to large-sized dogs to achieve an appropriate fit/seal.

The dogs in this study had normal lungs. How these results extrapolate to patients with pulmonary compromise remains to be determined.

Glucagon-like peptide-1 receptor agonists (GLP-1RAs) such as exenatide show promise in the treatment of feline diabetes. GLP-1RAs stimulate insulin secretion by pancreatic β cells in the presence of high glucose levels. The investigators in this study developed a delivery system that allowed the slow release of a stable GLP-1RA analog, [Gln28]exenatide. The study first validated the pharmacokinetics and pharmacodynamics of exenatide vs [Gln128]exenatide in cats, after which the conjugate compound consisting of [Gln28]exenatide bonded to hydrogel microspheres was evaluated. The plasma half-life of the SC administered microsphere-[Gln28]exenatide conjugate was ≈40 days as compared with 40 minutes with the injected free peptide. The investigators concluded that GLP-1RA in this formulation is suitable for once-monthly SC administration in cats.

Differences in pharmacologic activity and effect in reptiles as a result of injection location has been anecdotally observed and assumed, and mention has been made in the literature that some pharmaceuticals should be administered in the cranial body to avoid rapid clearance or toxic renal concentration via the renal portal or hepatic portal systems.^1,2 This is the first published report that confirms anesthetic agents can have differing effects based on location of injection.

Alfaxalone is a lipophilic neuroactive steroid that acts as a potent γ-aminobutyric acid agonist. It has been found to have reliable anesthetic results in a number of nontraditional species, including herptiles,^3-6 and is primarily cleared via cytochrome P-450 metabolism in the liver and CNS. This study, in contrast to the previously mentioned studies,^1,2 evaluated the differences in depth and duration of anesthesia observed in snakes that were injected in different regions of the body. Snakes that were injected in the cranial third of the body had an overall deeper plane of anesthesia and for a longer duration as compared with snakes that were injected in the caudal third.

This difference is most likely a result of the first-pass effect of the agent through the hepatic portal system when injected caudally, confirming that the anatomy of these species can affect pharmacologic effects.

Key pearls to put into practice:

A functional knowledge of anatomic differences is critical to high-quality, effective herptile medicine.

Specific tissue clearance of pharmacologic agents should be considered when selecting drugs, dosages, and administration route and location in reptiles.

Anatomy and physiology must be considered in all cases of nontraditional species medicine and therapeutics.

Possible causes of feline keratitis include viral infection (ie, feline herpesvirus-1), eyelid abnormalities (eg, eyelid coloboma, agenesis, entropion, distichiasis, ectopic cilia, tumors), ocular trauma, ocular foreign bodies, corneal sequestra, and bacterial infection. Although bacterial keratitis is less common in cats as compared with other small animals, clinicians should consider the role bacteria can play in keratitis and know how to identify and effectively treat corneal bacterial infections.

This retrospective study of 81 cats (102 corneal samples) describes clinical characteristics of cats diagnosed with feline keratitis, as well as in vitro susceptibility patterns of corneal bacterial isolates. Most patients were presented with unilateral disease and exhibited blepharospasm and ocular discharge.

Gram-positive bacteria were most often cultured (82 out of 102 samples), with Staphylococcus spp isolated from 55% of samples. Most samples (62 out of 81 cats) contained a single bacterial isolate. All isolates were susceptible to ofloxacin; other effective in vitro antibiotics included ciprofloxacin, ticarcillin, gentamicin, and moxifloxacin. Although chloramphenicol and doxycycline were effective in vitro, the authors did not recommend them as first-line therapeutics due to their bacteriostatic activity. Overall success for maintaining vision and globe retention was very good (88%) in this study.

Key pearls to put into practice:

Bacterial keratitis is uncommon in cats. When present, gram-positive bacteria, particularly Staphylococcus spp, are the most likely infectious agents.

First-line antibiotics to treat suspected bacterial keratitis include ofloxacin, ciprofloxacin, ticarcillin, gentamicin, and moxifloxacin.

Judicial empiric use of antimicrobials is essential to prevent antibacterial resistance. Culture and susceptibility testing is recommended when possible.

French bulldogs, English bulldogs, and pugs can be grouped together based on brachycephalic anatomy. They are known to have respiratory difficulty due to shortened noses and share a propensity for congenital vertebral malformations. These breeds are also known to have block vertebrae, hemivertebrae, transitional vertebrae, and neural tube defects.¹ A study revealed that the degree of screw-tail in French bulldogs correlates with the severity of hemivertebrae in the thoracic region.²

The current study examined 149 CT scans of vertebrae L6 to S3 and coccygeal vertebrae in neurologically normal pugs, French bulldogs, and English bulldogs over a 6-year period. The goal of the study was to determine whether vertebral lumbosacral (LS) malformations were present in neurologically normal dogs and whether the severity of tail deformity was linked to the presence of vertebral malformations in the LS region. Fifty-one percent of dogs had evidence of at least one type of congenital vertebral malformation, 60.5% had LS intervertebral disk herniations, and 67.1% had abnormal tails; normal tail morphology was only identified in 32.9% of dogs. These results support an association between LS hemivertebrae at L7 and S1 and the degree of tail malformation and intervertebral disk herniation in English and French bulldogs. Tails were more consistently normal in pugs, and this breed exhibited more transitional LS vertebral malformations than hemivertebral malformations. All breeds had an increased incidence of intervertebral disk disease as age increased.

This study concluded that the severity of screw-tail in English and French bulldogs is correlated with the presence of hemivertebrae; pugs do not have true screw-tails and are more likely to have transitional vertebrae. In addition, apparently clinically normal pugs and English and French bulldogs can have vertebral abnormalities. Because these results were found in clinically normal dogs, the study authors caution against overinterpreting results of CT scans and emphasize the importance of lesion localization during neurologic examination to avoid intervening in a clinically normal patient. In addition, there were some limitations due to the study’s retrospective nature and the fact that most dogs did not undergo neurologic examination; it is possible that patients with intermittent or mild neurologic deficits were considered normal.

Key pearls to put into practice:

A genetic defect of the DVL2 gene that is linked to vertebral malformations of the thoracic and coccygeal vertebrae has been identified in English and French bulldogs.³ Pugs do not share this defect and therefore should not be considered to have true screw-tails. The tail defect in pugs is more likely vertebra curva, the result of bone bending from soft tissue tension during development.

The study findings further support minimizing severe screw-tails due to their correlation with vertebral malformations. Although patients may be clinically and neurologically normal, selective breeding to minimize the screw tail phenotype may improve the gene pool.

Although advanced imaging can be helpful for visualizing anatomic neurologic lesions, the physical examination, including a thorough neurologic examination, is the most important factor in determining the relevance of imaging findings. In some cases, findings could be consistent with normal aging and not be clinically problematic.

This retrospective study of 371 cats treated for dermatophytosis sought to determine how often a first negative fungal culture was indicative of mycological cure as compared with 2 negative fungal cultures. Results demonstrated that a first negative fungal culture for Microsporum canis indicated mycological cure in 90.3% of cats, and subsequent cultures remained negative. Of the remaining cats, other than being lesional, 19 were healthy and had 1 negative fungal culture within the first 3 weeks of treatment, followed by ≥1 positive fungal cultures; these cats went on to mycological cure without event. Another 17 cats had concurrent medical illness in addition to dermatophytosis. These patients initially had resolution of lesions and negative fungal cultures but ≥1 positive fungal culture during treatment; mycological cure was not achieved until the concurrent illness was resolved. In otherwise healthy cats in which high treatment compliance is achieved, 1 negative fungal culture may be sufficient to indicate mycological cure.

AD can impact patient and owner quality of life.⁵ Signs of AD include pruritus, erythema, and development of other skin lesions and secondary skin infections.^1,2 The most common body regions affected include the axillae, inguinal area, interdigital spaces, muzzle, concave pinnae, antebrachial fossae, and periocular areas.⁶

Self-trauma and skin infections can exacerbate inflammation.³ Secondary skin and ear infections from Staphylococcus spp and/or Malassezia spp are common and tend to recur, which exacerbates the damaging cycle in the skin that perpetuates the condition.⁷ This propensity for relapse, as well as AD’s attribution to genetic and environmental factors, makes AD similar to its counterpart eczema in human medicine.^8-11

AD is thought to be caused by an immunoglobulin E (IgE)-mediated hypersensitivity immune response; these allergy response-associated antibodies are found mostly in the skin, mucous membranes, and lungs. AD has traditionally been thought to be caused by a response to inhaled allergens (eg, pollen, dust, dander, mold)²; however, newer evidence in dogs suggests that the cutaneous route of exposure to allergens is the most important.¹⁰ The underlying cause of AD is not fully understood,¹  but increasing evidence shows that skin barrier dysfunction is prevalent in atopic dogs.⁶  It is unknown whether the skin barrier defect is a primary or secondary issue.¹² Evidence suggests that inflammation disrupts skin barrier properties and likely contributes to a vicious cycle in dogs with AD in which an abnormal skin barrier increases the propensity toward allergen sensitization and exposure and then skin barrier function is worsened further by allergic inflammation response.^6,7,13 These issues can also lead to secondary bacterial and yeast skin infections, which further exacerbate the cycle.^1,2 Common components of skin barrier dysfunction include skin dysbiosis, structural changes, and altered intercellular lipid content, all of which allow for transepidermal water loss (TEWL) and penetration by allergens.^7,9 Studies have shown that TEWL is greater in atopic dogs than in normal dogs, increases with allergen exposure in atopic dogs, is greater in younger atopic dogs than in adult atopic dogs after exposure, and is lower during remission as compared with atopic dogs prior to treatment.^10,14 Allergen penetration activates an immune response that exacerbates inflammation and clinical signs primarily through degranulation of histamine-releasing mast cells, although other inflammatory mediators and immune cells are also involved.³

Altered intercellular lipids contribute to increased TEWL and penetration by allergens.¹⁵ In addition to free fatty acids, cutaneous ceramides are a major lipid component of the stratum corneum (ie, outer skin layer).^15,16 As compared with normal dogs, dogs with AD have decreased levels of both free fatty acids and cutaneous ceramides in skin lesions and skin that looks grossly normal.¹³ In a study, after being challenged with allergens, ceramide levels in atopic dogs returned to prechallenge levels within 2 months of lesion remission.¹⁷

AD is usually managed through various therapy combinations. A number of products, including Janus kinase inhibitors, monoclonal antibodies, and corticosteroids or immunosuppressants, are traditionally used in the management of canine AD. Allergen-specific immunotherapy is the only treatment option that may lead to allergen tolerance over time¹⁸; although a safe option, allergenspecific immunotherapy can be costly and take 6 to 9 months for initial observations of efficacy.¹

Other options for AD may be costprohibitive to the pet owner, cause adverse effects, and/or be incompletely effective alone.^1,19 Topicals and antihistamines can be less expensive but tend to have lower efficacy and duration of effect.^1,2,20 Therefore, research into additional supplemental management options for AD could be beneficial. 

Extracts derived from hardy kiwi (Actinidia arguta) fruit have been shown to change Canine Atopic Dermatitis Extent and Severity Index (CADESI)-03 scores in atopic dogs when used with steroids. CADESI is a subjective scoring system used to assess the severity of AD and monitor the condition over time, especially in clinical trials.²¹ Hardy kiwi extract has also been used to maintain lower CADESI scores after steroids have been tapered off.¹

This maintenance is likely achieved through the effects of the extract shown in regulation of allergic response and inflammatory mediators.^1,11,22-28 The fruit contains phenolic acids (ie, quinic and citric acid), monoterpenes, and flavonoids that may contribute to the activity of its extracts.²⁷ The duration of administration has been correlated positively with the effects and benefits. Hardy kiwi extract is best used long-term as part of a multimodal approach to therapy.^1,22

Ceramides are commonly considered for topical application, but evidence for oral administration is emerging.^12,29-32 Ceramides present in the skin represent 35% to 40% of the intercellular cement that binds the cells of the stratum corneum to provide an effective skin barrier.¹² Positive effects with oral ceramides in humans have been demonstrated on skin hydration and TEWL, IgE reduction, and reduction of other inflammatory mediators.^12,29-33

β glucans are found in nature as a structural component of the cell wall of yeast, bacteria, and fungi.³⁴ They are known for their immune-modulating effects and may help activate both innate and adaptive responses involved in AD.^34,35 In a double-blinded, placebo-controlled study in dogs with AD, mean overall improvement of pruritus, erythema, scaling, and lichenification was 63%.³⁶ These effects are likely associated with the modulation of inflammatory mediators.³⁷

Because of their safety and low cost, essential fatty acids are a common component of a multimodal approach to managing canine AD.^1,8 Inflammatory mediator, pruritus, and drug-sparing benefits have been demonstrated.^38-40 Linoleic acid is commonly used, as it is the precursor to γ linolenic acid; however, due to the absence of appropriate enzymes, this conversion is not possible in the skin and overall may be systemically deficient in atopic patients.^3,20,40-42 Providing performed long-chain omega-3 fatty acids and γ linolenic acid has had effects on TEWL, pruritus, IgE levels, inhibition of histamine release, and modulation of inflammatory mediators involved in AD.^3,20,40-47

Canine AD is a chronic condition that can be difficult to manage and frustrating for owners and veterinarians alike. Traditional approaches may be only partially effective alone, are sometimes cost-prohibitive, and can potentially cause adverse effects. Supplemental options, including hardy kiwi extract, ceramides, β glucans, and essential fatty acids, such as Dermaquin, provide an opportunity to support the efforts of a multimodal approach, potentially reduce pharmaceutical use—and therefore possible reduction of adverse effects—and promote maintenance of a healthy skin barrier in a cost-effective manner.

When a patient is presented for red and/or bruised ocular tissue, there may be concern for trauma or nonaccidental injury/physical abuse. In a large study of pets presented for traumatic injury, subconjunctival hemorrhage (ie, bleeding between the conjunctiva and sclera) was associated with nonaccidental injury.¹

However, there are several nontraumatic causes of subconjunctival hemorrhage that must be ruled out to avoid missing a systemic problem. These causes, in addition to cases of known trauma, were investigated in a recent study. Medical records of 147 dogs with subconjunctival or scleral hemorrhage were retrospectively analyzed. In 81% of dogs, subconjunctival hemorrhage was attributed to a traumatic event (eg, vehicular trauma, animal attack); of these cases, <5% were the result of nonaccidental injury. The remaining 19% of patients were determined to have a primary systemic or ocular problem that led to subconjunctival bleeding. Because a significant number of patients in this study had distinct systemic causes for subconjunctival hemorrhage, it is important to consider diagnoses other than trauma or nonaccidental injury.

Key pearls to put into practice:

When patients with subconjunctival hemorrhage are assessed, a thorough history, including the potential for unwitnessed trauma, should be obtained. A complete physical examination is necessary to look for evidence of puncture or bite wounds or abrasions. The patient should also be examined closely for any signs of petechiation or ecchymoses. A thorough ocular examination should be performed to assess for other signs of bleeding (eg, hyphema, iridal hemorrhage, retinal hemorrhage). Presence of additional intraocular signs may indicate a systemic problem or primary ocular condition (eg, glaucoma, orbital mass).

Diagnostic testing is important in cases in which trauma is not strongly suspected or observed. Blood pressure >160 mm Hg may be a concern for systemic hypertension, but pain, stress, and anxiety following trauma can also cause transient elevation in blood pressure. CBC, including platelet count, and prothrombin and activated partial thromboplastin times (aPTT) should be performed to rule out coagulopathies such as immune-mediated thrombocytopenia and rodenticide toxicity (Figure). A serum chemistry profile is also recommended to look for bleeding disorders secondary to acute liver injury or toxicity. In some cases, vasculitis secondary to rickettsial disease, envenomation, and/or another inflammatory condition may lead to a bleeding disorder.

Although nonaccidental injury is possible, subconjunctival hemorrhage may signal a bleeding disorder rather than abuse or trauma.

Cystocentesis (ie, obtaining urine directly from the urinary bladder by inserting a needle through the body wall) is common and considered the ideal method for obtaining urine for urinalysis and culture and susceptibility testing.¹ Cystocentesis typically helps prevent contamination (eg, from bacteria, RBCs, WBCs, or debris from the lower urinary tract and perineum) that can occur with voided or catheterized urine samples.² Diagnosis of bacterial cystitis is somewhat simplified through cystocentesis, as samples through this method should be sterile³; quantitative bacterial counts should be analyzed when evaluating voided samples.⁴ Although frequently iatrogenic and self-limiting, hematuria may be noted in cystocentesis samples.⁵

The urinary bladder is located in the caudal abdomen and positioned within the pelvic inlet ventral to the colon. The urinary bladder may fall to the dependent portion of the caudal abdomen if the patient is in lateral recumbency. The ureters terminate in the dorsal aspect of the bladder at the trigone. The urinary bladder is primarily composed of striated muscle, and blood is supplied by cranial and caudal vesical arteries. When filled, the urinary bladder can be easily palpated in most patients; however, palpation may not be feasible in some obese patients or in patients with either a low volume of urine or anatomic abnormalities (eg, pelvic bladder).⁶

Indications for cystocentesis include collecting urine samples from patients for which a urine sample is needed. Most cystocentesis samples are used for urinalysis and urine culture and susceptibility testing. Other diagnostic tests may include urine protein:creatinine ratio, urine cortisol concentration, urine catecholamine concentration, urine leptospirosis PCR, and urine electrolyte concentration.

Cystocentesis may also be used therapeutically in patients that have urinary obstruction secondary to uroliths, urethral plugs, and/or neoplasia. Decompressive cystocentesis can alleviate patient discomfort prior to urinary obstruction removal and can lower intraluminal bladder pressure and facilitate retropulsion of urethral plugs/uroliths, potentially easing catheterization.⁷ Decompressive cystocentesis should be performed with an extension set and a 3-way stopcock to allow for a single needle insertion as opposed to multiple needle insertions. Although decompressive cystocentesis has previously been discouraged, studies have suggested there is minimal risk for bladder rupture/uroperitoneum.⁸

Cystocentesis is often performed with ultrasonographic guidance, although this is not required (ie, blind cystocentesis). Using ultrasonography can help direct visualization of the needle in the urinary bladder lumen, avoiding iatrogenic damage of surrounding structures (Figure 1). The relative size of the bladder, echogenicity of its contents, and any obvious structural abnormalities can also be observed. A primary advantage of blind cystocentesis is that it does not require special equipment. Cystocentesis can be challenging to perform in patients that have abdominal effusion and should be performed with ultrasonographic guidance in such cases.

Cystocentesis is usually performed in awake patients, but sedation should be considered in fractious or uncooperative patients. See Related Article for a full outline of a sedation protocol.

Blind cystocentesis is contraindicated in patients with reproductive disorders (eg, pyometra); use of ultrasonographic guidance is recommended for such patients. There are no specific contraindications to blind cystocentesis in male dogs, although blood vessels on either side of the prepuce should be avoided (Figure 2).

There are some conditions, however, in which cystocentesis should be avoided entirely. Cystocentesis should be avoided in patients with pyoderma to prevent possible introduction of bacteria into the abdominal cavity and in patients that are unstable, as putting unstable patients in dorsal or lateral recumbency may lead to decompensation of hemodynamic status. Cystocentesis is also contraindicated in patients that may have bladder neoplasia, as cystocentesis in these patients can increase the risk for seeding neoplastic cells into the abdominal cavity.⁹

Coagulation status (ie, prothrombin time, activated partial thromboplastin time, thromboelastography, platelet count) should be considered before performing cystocentesis in patients at risk for bleeding. In addition, laceration or perforation of intra-abdominal blood vessels (eg, aorta) can lead to life-threatening hemorrhage.¹⁰ Abdominal exploratory surgery may be required in patients that develop hemoabdomen or uroabdomen following cystocentesis that cannot be stabilized with conservative management.^10,11

Other potential complications include bladder wall hematoma, bladder wall rupture leading to uroabdomen, intravesicular blood clots, focal peritonitis, peritoneal free gas, abdominal wall abscessation, nodular fat necrosis, aortic or bladder wall hematoma, and bladder mucosal detachment.¹² Vasovagal collapse has also been reported in cats following cystocentesis.¹³ In addition, one case report describes a dog with bacterial cystitis that developed septic peritonitis following cystocentesis.¹¹

After cystocentesis, a small amount of gas may be introduced into the urinary bladder iatrogenically but is typically of no clinical consequence (Figure 3). Evidence of this can be seen on ultrasonography after the procedure is completed and appears as horizontally oriented, parallel hyperechoic lines reverberating off the introduced gas.

Depending on clinician preference and patient temperament, a staff member may hold the patient. Patients can be positioned in either dorsal or lateral recumbency. The authors prefer lateral recumbency in cats. In female dogs in dorsal recumbency, the bladder is usually located below the umbilicus, where isopropyl alcohol pools. A V-trough may help maintain patient comfort in dorsal recumbency. For patients in dorsal recumbency, the urinary bladder is often located on the midline between the fourth and fifth nipples. In male dogs, the prepuce makes aspiration of urine on the midline difficult. Paramedian insertion of the needle is acceptable; however, the caudal superficial epigastric veins that lie on either side of the prepuce should be avoided. Standing cystocentesis, which involves puncturing through the lateral abdominal wall, has been described as safe and effective and may be less stressful for patients.¹⁴

Although there are no evidence-based recommendations, the authors recommend preparing the ventral abdomen by clipping the hair and scrubbing the proposed insertion site to minimize the risk for iatrogenic bacterial peritonitis (Figure 4).

• Clippers and antiseptic solution (dilute chlorhexidine or iodine-based preparation)
• 1- to 1.5-inch, 22-g needle
• 3- to 12-mL syringe
• Collection tubes (glass red top tube or any empty tube without preservatives).
  • A nonadditive sterile tube is generally recommended for culture. There are other tubes available that may aid in keeping bacteria viable in transit. A reference laboratory can typically help identify appropriate tubes for urine-based tests.
• Ultrasound machine (for ultrasound-guided cystocentesis only)
• Sedating agent, if needed
• 70% isopropyl alcohol
• Additional supplies if removing a large volume of urine
  • Extension set
    • Alternatively, a butterfly catheter can be used in smaller patients in lieu of an extension set and needle
  • 3-way stopcock
  • Larger syringe (12-60 mL)
  • V-trough (optional)

Place the patient in either dorsal or lateral recumbency. If the patient is a male dog, gently retract the penis off the midline.

Gently palpate the caudal abdomen to isolate and secure the urinary bladder.

Using an ultrasound probe, visualize the urinary bladder. Insert the needle attached to a syringe roughly 5 to 10 mm cranial to the ultrasound probe. Introduce the needle at a 45° angle to the transducer probe and pass it into the abdomen in a cranial-to-caudal orientation.

Visualize the needle (appears as a hyperechoic line with distal shadowing [seen in the top left of the Figure]), then advance it into the urinary bladder lumen, terminating near the junction of the bladder and urethra (seen on the right side of the Figure).

Apply negative pressure to the syringe to obtain the sample, then release the negative pressure and slowly withdraw the needle.

With the patient in either lateral or dorsal recumbency and a staff member holding the patient, gently palpate the caudal abdomen and isolate the urinary bladder.

Using gentle digital pressure, secure the urinary bladder with one hand. If the patient is in dorsal recumbency (Figure), insert the needle with the syringe attached in a cranial-to-caudal orientation and a 45° angle to the body wall. If the patient is in lateral recumbency, insert the needle in a cranial-to-caudal orientation while approaching the urinary bladder from its lateral aspect.

Apply negative pressure to the syringe to obtain the sample, then release the negative pressure and slowly withdraw the needle.