When news first broke that SARS-CoV-2, the virus that causes COVID-19, was believed to have originated from bats, questions quickly surfaced about the potential threat of infection from companion animals to humans and vice versa. Initially, veterinary experts found no evidence of a correlation. However, that changed when animals 

 and then in the U.S. tested positive for the virus. The first reported case in the U.S. was a tiger at the 

As human medical experts raced to identify the chain of transmission and possible cures, their counterparts in the veterinary field searched for links on how the virus could be transmitted through human-animal contact and what impact that might have on animal patients.

“We were really interested at the outset in how prevalent this spillover might be,” explained Jonathan Runstadler, DVM, PhD, a professor of infectious disease and global health at Cummings School of Veterinary Medicine at Tufts University, during a recent webinar hosted by the Massachusetts Veterinary Medical Association. “Part of the problem initially was that in late January and through February, there wasn’t information or data of any sort to look at in order to evaluate if this was a risk.”

Dr. Runstadler provided an overview of the university’s Coronavirus Epidemiological Research & Surveillance (CoVERS) study. “The study we’ve started grew from some of what we already know about viruses like SARS-CoV-2 and related viruses that have caused epidemic events in the past,” he said.

“We know that these viruses are spilling over into human populations from animal hosts. It is usually animal hosts in the wild but [it also occurs] through domestic intermediate hosts or other animals that are in semi-domestic situations. For SARS-CoV-2, that really is no different.”

Quickly shifting focus from the department’s ongoing studies, the research team started locally by sampling patients from veterinary hospitals in Massachusetts. Initially, they hoped to determine whether significant spillover was occurring in situations where close human-animal contact was common. As the study’s focus began to take shape, however, the CoVERS team established the following three key phases:

In the first phase, they screened a variety of companion and large animal species to estimate the transmission of SARS-Cov-2 to animal hosts. The subjects were chosen without regard to the COVID-19 status of owners or other humans with which the animals had contact.

In the second phase (where the team is currently focused), the investigators are screening and monitoring paired owners and animals to evaluate direct SARS-Cov-2 transmission and identify high-risk situations.

The third phase will focus on monitoring farm and veterinary school animals that are in regular contact with humans for natural SARS-CoV-2 infection as well as inter- and intraspecies transmission over the course of the COVID-19 pandemic.

The first animal to be tested for the CoVERS project was a hedgehog. The group has also tested horses, bats, a coyote and even seals, but Dr. Runstadler said most of the samples have been taken from dogs and cats.

“Based on some of what we know from experimental inoculations of animal hosts and from what has occurred naturally as well, it appears that cats may be the most interesting population in terms of domestic animals.”

Of the more than 350 animals that have been tested through CoVERS to date, there have been no cases of SARS-CoV-2. “We are glad that we have not detected positives in the population of patients that we’re looking at,” Dr. Runstadler said.

Looking ahead, the team is interested in enrolling more human-animal pairs for the home phase of the study, which they hope includes adding human samples for testing. “If we get a spillover event we will be able to look at the virus coming from the human as well as the spillover virus,” he explained. “That may be important for understanding how this virus actually does spill over, when it does in those rare instances, between humans and animals.”

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So far, none of the animals involved in the CoVERS study have tested positive for the virus that causes COVID-19. 


Study examines human-to-animal transmission of SARS-CoV-2 

High-risk neonatal puppies and kittens are those that become severely dehydrated or systemically ill, or are falling behind in weight gain. A variety of factors can contribute to these circumstances, of which husbandry is the most common. Failure of passive immunity and/or infectious disease can also contribute.

Caring for these high-risk neonates requires knowledge of normal newborn parameters. Healthy puppies and kittens should be gathered in a loose pile within their enclosure (

), should have good mucous membrane coloring and should be noted to be “twitching” while asleep; they should not be extremely vocal.

A healthy puppy or kitten will twitch noticeably, almost continuously, while sleeping (

). This can even startle them awake or cause them to readjust their sleeping position. In my experience, evaluating for this twitching has been the most important factor when monitoring neonates. A significant decrease or lack of twitching could signify that the animal is not doing well and requires professional evaluation.

Several other factors should be monitored as well when caring for high-risk neonates, including environment, hydration, nutrition, excrement, weight and immunity.

In the first week of life, puppies and kittens rely solely on their external environment for warmth; they are unable to maintain their own body temperature. Environmental temperatures ranging from 75°F to 80°F will support maintaining an optimal neonatal rectal temperature of 98°F to 99°F. Only when a puppy or kitten is maintained at an appropriate temperature is it able to digest milk and thus ready to be fed.

High-risk neonates are typically solitary individuals; an entire litter of high-risk neonates is rare. Those at high risk prefer to snuggle against something warm. IV fluid bags warmed in the microwave (make sure the bag is neither too hot nor too cold) and covered with a tube sock can provide a nice substitute for littermates. A sock will be more inviting to the neonate and, because they can be changed and laundered easily, socks also help keep the area clean.

A well-hydrated puppy or kitten will have adequate skin turgor and good pigment color. Light-colored animals should have bright pink mucous membranes (

). As for adults, it can be challenging to assess mucous membranes in darkly pigmented neonates. However, there should be areas where assessment is possible, such as the belly.

Any fluids administered to a neonate, regardless of route, should be warmed. For mildly dehydrated newborns, subcutaneous fluid administration is appropriate. Crystalloid fluids should be administered at a dose of 1 ml/30 g. In cases of severe dehydration, an intravenous or intraosseous catheter is greatly preferred. Jugular intravenous catheters may be the easiest to obtain, and the femur is an excellent bone for intraosseous placement using a hypodermic 22- or 20-gauge needle. Both routes can be used to administer fluids, blood products, diluted dextrose and antibiotics. Severely hypovolemic neonates can receive approximately 60 ml/kg of crystalloid fluid total during one resuscitation event. In cases of dehydration, the rehydration process should be more gradual.

Dehydration deficits in neonates are calculated in the same way as for adults: Percent dehydration multiplied by body weight in kg will provide the total fluid deficit in liters.

 The neonate’s maintenance fluid is about 60 to 100 ml/lb/day.

 Colloids can be used in urgent circumstances and are dosed the same as in adults.

Any species-specific milk replacer or goat’s milk can be used to feed neonatal puppies and kittens. Neonates need to be fed every two hours until they are gaining weight steadily and becoming more glossy in appearance. They can be bottle fed only if they are vigorous enough to perform the task adequately. Bottle feeding should not be attempted until the neonate’s Apgar score is between 7 and 10 (

 Neonates with Apgar scores between 4 and 7 should be tube fed via a red rubber catheter (size 8-10 Fr for canine neonates and 5 Fr for neonatal kittens.)

Correct tube length is imperative when tube feeding. The tube should be aligned from the mouth and extended down to the last rib. When placing the tube in the stomach, you will feel resistance as the tube passes the lower esophageal sphincter. Place the animal in sternal recumbency and insert the tube toward the back of the throat on the left side, advancing rather swiftly. A puppy or kitten that is alert enough to be fed will also fight you to some degree during tube placement. Holding the mouth closed and the neck extended while milk is instilled will help keep the tube in place.

It is very common for neonates to grunt and vocalize while being fed. Hearing this is a sign that the tube has been placed correctly. An animal that is too lethargic to swallow the tube and provide some resistance is not quite ready for enteral nutrition. While feeding the neonate, be sure to palpate the area below the lower rib on the left side of the body to ensure you are not overfeeding. A good guideline for feeding volume is approximately 4 to 5 ml/100 g. If the puppy or kitten is showing signs of discomfort during feeding, do not proceed with the entire volume but work up to it incrementally. To prevent residual milk from dripping into the pharynx, risking aspiration, kink the tube before removing it.

Because neonates cannot urinate or defecate without assistance, they should be stimulated both before and after feedings. To stimulate urination, rub the ventral caudal abdomen/inguinal area. Urine should be almost clear in a well-hydrated neonate (urine specific gravity < 1.017). A neonate with darker urine will need a more aggressive fluid therapy plan.

Some neonates may urinate and defecate simultaneously, while others need separate stimulation to produce feces. Do not rub directly on the anus repeatedly to stimulate defecation; this can result in ulceration of the anal tissue and make defecation uncomfortable. Instead, wipe a damp, warm cloth or gauze bilaterally on either the ischium or the pelvis (or both). If ulcerations do occur, use vitamin A & D ointment in between stimulation events.

Neonatal feces should resemble varieties of mustard. It can be bright yellow or darker yellow, with or without “seed” material in it. The seeds are fatty excrement. The feces can be very wet in appearance but should not be completely liquid. The meconium is much darker and gelatinous. At-risk neonates that have not had adequate nutrition may pass meconium much later after birth.

A healthy puppy or kitten will gain about 10% of its body weight daily. High-risk neonates should be weighed twice daily using a gram scale. This will allow minor weight losses and gains to be observed more closely to allow for a necessary intervention sooner rather than later.

Neonates that have failed to nurse on their dam or queen within 18 hours of birth will not have had the opportunity to ingest colostrum. In these cases, plasma or serum can be used as a substitute. If administered within the first 18 to 24 hours of life, the plasma or serum can be given orally to the neonatal puppy. Puppies should receive 0.1 ml/g of adult dog serum.

In kittens, care should be made to ensure the serum blood type matches that of the kitten. Dosing for kittens is 15 ml non-hemolyzed adult serum/100 g of body weight. This serum should be divided into three total injections given every eight hours for the first 24 hours of life.

Newborn puppies and kittens can become compromised quickly, particularly in the first week of life. Their owners may miss early warning signs of compromise, only noticing that something is amiss when the neonate has entered a high-risk state.

Dr. Simermeyer is an emergency practitioner at Mount Laurel Animal Hospital in Mount Laurel, NJ. Her professional interests include emergency and critical care, and canine reproduction. She has been breeding Cardigan Welsh Corgis for the past 23 years.

Thomovsky EJ. How to treat the sick neonate. Presented at the Atlantic Coast Veterinary Conference; 2015.

Veronesi MC, Panzani S, Faustini M, et al. An Apgar scoring system for routine assessment of newborn puppy viability and short-term survival prognosis. 

Newborn puppies and kittens can become compromised quickly. Here’s what you need to know when caring for these fragile animals.

Neonatal nursing care for high-risk puppies and kittens

Transitioning from undergraduate to veterinary school can be tough to say the least, largely because of the massive workload and fast-paced learning, says Danny Sack, DVM, a 2020 graduate and small animal intern at the Animal Medical Center in New York. One good way for first-years to stay afloat is to understand how they learn best, he tells dvm360.

Speak to your professors, speak to different clinicians that you get along well with, and ask what worked for them in veterinary school, says Dr. Sack. Also speak to your peers, and ask how they’re studying. "But take every piece of advice with a grain of salt," he advises. "Not everybody learns the same [way]," he says. 

He also advises first-year students not to compare themselves with others. “It’s really important to stay in your lane and do what makes you be the best student you can be, and that will kind of keep you on track and limit the stress you don’t need in your life,” he says.

Finally, he emphasizes the significance of proper self-care. His advice: Try not to push yourself past your limit and make time to rest.

“Don’t grind to the point where you can’t get up the next morning, where you’re having mental breakdowns because you can’t stand it one more minute," says Dr. Sack.

Watch the video below for more of Dr. Sack’s recommendations.

Videos

Danny Sack, DVM, offers some tips to help new veterinary students navigate their most challenging year of veterinary school.

How to survive your first year of veterinary school

During periods of immobilization and restricted activity, muscles atrophy as a result of increased myostatin. Myostatin is a protein that triggers muscle breakdown and hinders muscle protein synthesis and muscle cell differentiation.

 This phenomenon especially reveals itself during the requisite 8 to 12 weeks of postsurgical “down time” following repair of cranial cruciate ligament (CCL) rupture, the most common stifle injury in medium- and large-breed dogs.

The commercial supplement Fortetropin (MYOS Rens Technology), a freeze-dried, fertilized chicken egg yolk product, reduces myostatin. Human studies have shown that Fortetropin abates serum myostatin levels by 18% to 22% and increases muscle thickness and lean body mass; Fortetropin has similar effects on serum myostatin in rats.

 In both healthy dogs and golden retrievers with muscular dystrophy, reduction of serum myostatin boosts muscle mass.

 researchers at Kansas State University’s Veterinary Health Center investigated the effects of Fortetropin supplementation on disuse muscle atrophy. A total of 100 client-owned dogs with naturally occurring CCL ruptures were randomized into two groups: a Fortetropin treatment group and a placebo-control group.

All dogs in the study weighed between about 39 and 165 lb (18-75 kg); had a diagnosis of naturally occurring CCL rupture; had no surgical correction of the contralateral CCL within the six months prior to enrollment; and had no concurrent disease.

One-third of the enrolled dogs were purebred or mixed Labrador retrievers. Golden retrievers, boxers, Rottweilers, American bulldogs and American Staffordshire terriers were also highly represented. Seventy-one of the dogs were female and 29 were male; the average age was 5.5 years. Bilateral CCL ruptures were present in 16 dogs.

Each patient underwent tibial plateau leveling osteotomy (TPLO), the most common surgical repair technique for CCL rupture.

 The Fortetropin group, composed of 52 dogs, was dosed at 300 mg/kg of the powdered supplement mixed with canned food once daily for 12 weeks. The placebo group, made up of 48 dogs, received a cheese powder.

The following assessments were performed preoperatively on each dog, and then at 8 and 12 weeks postoperatively:

Thigh circumference, measured with a retractable tape measure at the midpoint of the femur for both the affected and unaffected legs

Muscle thickness, assessed by B-mode ultrasonography cranial and lateral to the mid-femur of both the affected and contralateral hindlimbs; and dorsal to ventral transverse thickness of the epaxial muscles at the level of the 13th rib for both the affected and unaffected sides

Stance analysis, or the percentage body weight placed on each limb at a static stance

Some dogs did not finish all evaluations. Complete data sets were obtained for 77 dogs for muscle measurements and stance analysis, and for 69 dogs for serum myostatin measurement.

Dogs that received Fortetropin experienced no significant change in thigh circumference for either hindlimb over any time periods, while dogs receiving placebo had a significant reduction from weeks 0 to 8. For ultrasound muscle measurements, however, there was a significant decrease in the Fortetropin group from weeks 0 to 8, but not from weeks 8 to 12 or weeks 0 to 12. Dogs given placebo had no significant changes in muscle thickness over any time periods.

Stance analysis for the Fortetropin group showed a 7.0% climb in weight supported by the affected hindlimb from weeks 0 to 12, compared with 4.9% for the placebo group. The corresponding drop in weight supported by the contralateral hindlimb from weeks 0 to 12 was 6.5% for the dogs receiving Fortetropin, compared with 5.8% for the placebo group.

Serum myostatin levels did not change significantly over any time period (0-8, 8-12 and 0-12 weeks) in the Fortetropin group. But in the placebo group, serum myostatin rose significantly from weeks 0 to 8, although not from weeks 8 to 12 or 0 to 12.

The addition of Fortetropin to the diet following TPLO surgery seemed to have a protective effect on thigh circumference compared with placebo, yet this benefit was not substantiated in the ultrasound measurements of muscle thickness. These measurements dropped significantly from weeks 0 to 8 in the Fortetropin group, and insignificantly in the placebo group. Investigators attributed this contradiction to inconsistency in the personnel performing the ultrasonographic measurements and to differing degrees of transducer force applied by the examiners. They also raised the possibility that thigh circumference measurements may have been impacted by variations in hair coat thickness and postoperative hair regrowth rates.

Nonetheless, compared with placebo, Fortetropin supplementation appeared to hasten weight-bearing capacity for the affected limb. Serum myostatin increased significantly in the placebo group during the period of exercise restriction but did not change significantly in the Fortetropin group. Preventing a rise in serum myostatin during periods of exercise restriction is key to curtailing disuse atrophy.

Fortetropin is an established agent for increasing lean body mass and cutting myostatin levels in people.

 But for dogs undergoing orthopedic surgery, the study described herein is the first published report on the protective effect of Fortetropin. When comparing Fortetropin with placebo administered during the postoperative exercise-restricted period, Fortetropin seems to prevent rise in myostatin and corresponding loss of muscle mass.

Dr. Capuzzi is a small animal veterinarian and journalist based in the Philadelphia area.

Elliott B, Renshaw D, Getting S, et al. The central role of myostatin in skeletal muscle and whole body homeostasis. 

Veterinary Surgery: Small Animal Expert Consult

Sharp MH, Lowery RP, Mobley CB, et al. The effects of Fortetropin supplementation on body composition, strength, and power in humans and mechanism of action in a rodent model. 

Freeman LM, Rush JE, Cunningham SM, et al. Pilot study of a myostatin antagonist in dogs with cardiac cachexia. 

5. White DA, Harkin KR, Roush JK, et al. Fortetropin inhibits disuse muscle atrophy in dogs after tibial plateau leveling osteotomy. 

6. Reif U, Hulse DA, Hauptman JG. Effect of tibial plateau leveling on stability of the canine cranial cruciate-deficient stifle joint: an in vitro study. 

The supplement Fortetropin has been shown to curb muscle atrophy associated with postoperative exercise restriction in dogs following surgical repair of cruciate tears.

Disuse muscle atrophy inhibited by nutritional supplement

With so many options available today, discussing 

 with your clients can be a complex and time-consuming affair. Some practices simply give clients a handful of brochures, while others research the available options themselves and recommend that clients select from among just two or three plans. In some cases, these tactics have left both veterinary practices and pet owners stuck with “analysis paralysis.”

But getting your patients insured doesn’t have to be difficult or frustrating. Here’s how to help your clients understand the benefits of pet insurance and enable you to deliver gold standard care.

The best time to discuss pet insurance is the very first time a client visits your practice with their pet. Many of these first-time patients are young and healthy, with no diagnosed pre-existing conditions. (No pet health insurance policy covers pre-existing conditions, so it’s important to get the pet insured before any illnesses appear.)

Discuss pet insurance in the broad context of the pet’s expected lifetime costs. Many pet owners need to be educated about potential health risks for pets and the true cost of pet ownership in order to understand the benefits of financial planning and protection.

Simply handing a client some brochures is not proactive education. Instead, turn this interaction into a conversation you have during routine care. Speak to your clients in the same way you might when discussing heartworm or flea and tick prevention: Taking simple and affordable action today can mitigate future costs. Keep in mind that veterinary practices 

 annually through insured pets than through uninsured pets.

Giving pet parents too many choices can actually reduce the chance that they will move forward and enroll in any type of coverage. However, veterinary practices should not take a one-size-fits-all approach when it comes to pet insurance providers either. The insurance discussion should be one of education and not necessarily specific recommendations. Educate your clients on the value and impact of insurance, then let them choose.

Some clinics opt to have a “pet insurance expert” in house who researches various insurance providers and then creates a detailed pamphlet for clients. Others leverage third-party online pet insurance comparison/education websites that do this research objectively and help guide clients through the insurance buying process.

Including these resources on your practice website will make it easy for clients to dive in on their own for more detailed information about pet insurance options. Also use social media to generate positive messaging about pet insurance. This is often a much more laid-back format for discussion, but those digital conversations can be as impactful as in-person interactions.

To make it easier to know which patients are insured, make note in the patient’s medical record. If they do have pet insurance, document the type of policy. This will help when making treatment recommendations, and it changes the conversation substantially.

For example, let’s say you need to recommend a $5,000 procedure. That’s a 

. But if your client has a good pet insurance plan and you can inform them that 80% of that cost will be reimbursed, they will be more likely to follow your recommendation.

Also, make note of what wellness benefits the client can allocate and inform them during their visit. This will help them use those benefits before they run out at the end of the year.

pet owners are more likely to buy pet insurance

 if their veterinary hospital’s team members have it themselves. If your staff members don’t use pet insurance, what signal does that send about pet insurance to your clients?

Some of your staff may provide treatment to their animals free of charge or at a discount, and feel that they don’t need pet insurance. However, you must keep risk and liability in mind. Recent IRS regulations state that veterinary practices cannot offer staff discounts of larger than 20% on services. Rather, IRS recommends that practices offer products and services at cost, plus 10%.

Dr. Hodges is an emergency relief veterinarian and advisor for 

, the only pet insurance marketplace actively endorsed by veterinarians. He has seen the positive 

 in emergency cases and believes removing financial barriers from important pet care decisions is a key step forward.

Help your clients avoid “analysis paralysis” when it comes to choosing a pet insurance plan.

How to frame the pet health insurance conversation with clients

“You know how you can walk into a restaurant and think, ‘ew, this place is gross,’ but then the service and the food are fantastic, so you keep coming back?” asks Jonathan Kaufman, DVM, owner of Eastern Animal Hospital in Baltimore, Maryland. “That’s how my old facility was—not much to look at, but our doctor and staff care and service were great, so our clients kept returning.” 

Precisely because client and staff care is so important, Dr. Kaufman knew his deserved a nicer facility. He had lived for eight years with the first practice he bought, then remodeled a shopping center site in 1998. Although he did the best he could on the remodel—completed with the help of architect Wayne Usiak of BDA Architecture—the space had its limitations. 

“It was clear to me that we really couldn’t grow anymore in that small space,” says Dr. Kaufman. “Demand was high, and we had nowhere to go. We had up to eight doctors and about 50 people all working in a 5,000-square-foot building. We were still doing a great job, but we were stressed out in such a small space.” 

Throughout this time, Dr. Kaufman had been talking to Usiak again about wanting to build a larger practice, this time designing from scratch. He was excited about the raw possibilities a new build would offer. When a two-acre lot of land became available, he jumped, knowing it was time to build the practice of his dreams. 

“Back to the restaurant analogy, it’s like our building is the` table on which we serve our clients,” he says. “A larger, nicer ‘table’ would allow us to serve them in a much more enjoyable way.” 

He built that table and earned Eastern Animal Hospital the title of runner-up in the 2020 dvm360 Hospital Design Competition. The 16,433-square-foot practice garnered praise from the Hospital Design Competition judges for its great traffic flow as well as a gorgeous lobby. 

Twenty years after his first remodeling experience, Dr. Kaufman said it was time to “go big or go home.” So big he went.

“My dream was to start with a raw piece of land and build an inspiring facility that would take our hospital into the future,” says Dr. Kaufman. “I wanted my staff members to have a building that was quiet and spacious with a clinical floor and an upper floor for administrative tasks, meetings and an eating area so they could get away from the intensity of clinical practice.” 

First on the list was designing a lobby with a smooth flow that sets the stage for the rest of the practice. Gone are the days of big, cavernous waiting areas, he says. Instead, Dr. Kaufman chose a design that allows for greeting the client, then quickly transitioning them into an exam room, then back out a different door after the exam or treatment, with checkout in the exam room itself for privacy. 

“Our emphasis was on the flow of the building, making a seamless transition from space to space and not making clients wait too long,” he says. 

The centerpiece of the lobby is a 1,000-gallon cylindrical saltwater tank, something he has had in every practice since the beginning. This one, however, is much grander than before. He also chose partitioned seating surrounding the tank in an effort to reduce pet/client anxieties, including being able to separate dogs and cats. The room features plenty of charging ports, internet access, bench seating for both owners and pets, and a client concierge center with a custom grinding coffee station, water and fresh snacks. 

Once through the lobby, clients are taken into one of 14 exam rooms, a big upgrade from the previous six. Here, Dr. Kaufman invites clients to “linger at the table.” 

“The idea behind not making clients wait too long in the lobby is like that of a restaurant: When you’re waiting for your table, you’re impatient, wondering what’s taking so long,” he says. “But once at your table, you want to slow down and take your time, with no pressure to clear the table for the next person. Having ample exam rooms allows us to slow down, not rush clients back out the door.” 

While the lobby and the gigantic fish tank are striking, Dr. Kaufman says the surgery suite is his favorite spot in the hospital. “The surgery room was designed with lots of windows, plenty of space, and glass looking out,” he says. “Surgery is what I love, and this space provides me with a very peaceful Zen feeling. It’s a beautiful room that lets me do my thing and do it well.” 

While a large, flowing hospital with the latest and greatest equipment was at the top of Dr. Kaufman’s list, creating a strikingly beautiful building came next. With the help of the experts at BDA Architecture, he got what he wanted. A porte cochère offers clients a covered entryway to the building, making an instant statement upon arrival. Upon entry, clients are greeted with a rustic stacked stone wall behind reception, modern lighting and other high-end features. 

“It feels like you’re checking into a high-end hotel, not a veterinary practice,” he says. “All the finishes are durable and beautiful, making for a truly lovely experience. In the end, the two-plus years it took to design and build the new Eastern Animal Hospital allowed me to bring my dream vision to reality.”

Sarah A. Moser is a freelance writer in Lenexa, KS.

Topnotch veterinary care deserves a topnotch facility to match.