Use of a vascular access port for antibiotic administration in the treatment of pododermatitis in a chicken.
Key words: multidrug resistant E coli, pododermatitis, vascular access port, portocath, ceftazidime, avian, chicken
A 4-year-old, male, Institut de Selection Animale (ISA) brown chicken was evaluated for treatment of chronic pododermatitis involving one foot. The bird had been rescued by the owners from a roadside 3 years earlier and was living in a large pen with the opportunity for free-range grazing. It was fed a mixed diet of a poultry mash, grain, mixed vegetables, and free-range food items.
Seventeen months before presentation, the bird had been diagnosed with mild pododermatitis and treated, at first conservatively but later surgically, at 2 different veterinary clinics. A variety of antibiotics had been used, including trimethoprim-sulfadiazine, enrofloxacin, and amoxicillin-clavulanic acid. Several attempts at curettage and bandaging, as well as placing doxycycline-impregnated polymethylmethacrylate beads, had been unsuccessful in effecting a good outcome. Results of a bacterial culture collected midway through this treatment regime had grown Klebsiella species, Escherichia coli, and an unidentified anaerobic bacterium. Both the Klebsiella species and E coli were sensitive to amoxicillin-clavulanic acid, but the E coli demonstrated resistance to many other antibiotics.
On examination the bird was in good body condition (weight 3.5 kg) with no obvious physical problems other than the right foot. The plantar surface of this foot had a deep ulceration, approximately 1.5 cm in diameter, with swelling of the surrounding tissues. The foot was hot and painful to the touch, and the bird exhibited moderate lameness. With the chicken under anesthesia, a deep swab sample of the wound was collected and the foot was radiographed. On radiographs, mild-moderate degenerative joint disease was visible in the right tarsometatarsal and phalangeal joints. The owner was instructed to flush the wound and change the dressing once daily. Oral meloxicam (0.3 mg/kg PO q12h) and tramadol (10 mg/kg PO q12h) were given for analgesia.
Results of an aerobic bacterial culture were a heavy growth of a multidrug resistant E coli bacterium, sensitive only to amikacin, imipenem, and third-generation cephalosporins (ceftazidime, cefoxitin, and cefotaxime). Although the cephalosporin antibiotics can be given by intramuscular injection, the anticipated duration of the treatment (several weeks or months), the frequency of the injections, and the volume of the drug to be injected raised animal welfare concerns. Accordingly, the decision was made to implant a vascular access port (VAP; Le Petit Companion Port, Norfolk Vet Products, Skokie, IL, USA) to facilitate intravenous administration of the drug, obviating these concerns.
The bird was premedicated with midazolam (0.2 mg/kg IM, Hypnovel 5 mg/mL, Roche, NSW, Australia) and morphine (1 mg/kg IM, DBL Morphine Sulphate Injection BB 5 mg/mL, Hameln Pharmaceuticals, GMBH, Germany), and then anesthetized by mask induction, followed by intubation and maintenance with isoflurane and oxygen. The right lateral cervical apteryla was surgically prepared at the base of the neck. A longitudinal incision was made approximately 10 cm from the base of the neck, over the jugular vein. The vein was then identified and blunt dissected from surrounding tissues for a length of approximately 3 cm (Fig 1). Suture loops (3-0 polydioxanone suture, PDS, Ethicon, Somerville, NJ, USA) were placed at both ends of the mobilized vein, and the incision was covered with a moistened swab. A second incision was made caudal and slightly dorsal to the first incision, at the base of the neck. A subcutaneous pocket, approximately 6 cm in diameter, was created by blunt dissection, and a tunnel was created between the 2 incisions (Fig 2).
A 4-Fr rounded-tip silicone catheter was passed through this tunnel, and the port end was occluded with hemostatic clamp. The free end (rounded tip) was then introduced into the jugular vein, directed toward the heart, via a peel-away needle introducer (Fig 3). The catheter had been premeasured so that the tip was positioned at the entrance to the right atrium. The VAP was then flushed and preloaded with 0.9% saline with a 22-gauge Huber needle. The occluded end of the catheter was taken through the subcutaneous tunnel, excess length was trimmed, and the catheter was attached to the barbed outlet pin on the VAP and secured. The VAP was flushed with 0.9% saline again with a Huber needle to confirm patency of the entire system, blood was aspirated, and the VAP was then flushed again with 0.9% saline. The VAP was positioned within the subcutaneous pocket such that the septum of the VAP did not lie under the skin incision line, and the VAP itself was not rotated within the pocket; it was then sutured to the underlying fascia with single, interrupted, 3-0 PDS sutures through each of the 4 anchor points on the base of the VAP (Fig 4). The subcuticular tissue and skin incisions were then closed in separate layers with 3-0 PDS in a continuous pattern (Fig 5). Patency was again confirmed as described above, and the VAP was "locked" with 2 mL of heparin-saline (100 units/mL). Anesthetic recovery was uneventful.
During the next 2 days, the chicken was hospitalized and received ceftazidime (20 mg/kg q12h, with a saline flush before and after the infusion, and the VAP was locked with 2 mL of heparin-saline after the second flush. No adverse effects were observed, and the bird was discharged to the owner after a training session to demonstrate the use of the VAP. The owner reported no difficulties in accessing the VAP and giving the antibiotics twice daily, and they cleaned the wound and changed the bandage each day.
After discussion with a microbiologist, it was decided to culture the wound every 2 weeks and continue treatment until 2 weeks after the third consecutive negative culture. Two weeks after commencing treatment, results of bacterial culture showed only a light growth of E coli; 1 month after commencing treatment, no growth was detected. Production of caseous exudate decreased and finally stopped during the next 2 weeks, and the skin over the plantar wound appeared healed after another month. Treatment was discontinued after 2 months, with the device flushed and locked twice weekly to maintain patency.
Follow-up examination 3 months after the wound had healed revealed that, although the plantar surface was healed, severe degenerative arthritis was present in many of the joints in the foot. This was being managed with tramadol (30 mg/kg PO q12h) and meloxicam (1 mg/kg PO q12h), and the bird appeared to be comfortable, maintaining its appetite and weight and behaving normally. With no evidence of infection, the decision was made to remove the VAP.
Under general anesthesia (as described above), the skin over the VAP was incised to expose the VAP and allow blunt dissection to free it from the surrounding fascia. The catheter was disconnected from the VAP, and a second incision was made over the jugular vein to allow the catheter (now encased in connective tissue "pseudotunnel") to be traced back to its entry into the jugular vein. This entry point was occluded with digital pressure while the catheter was removed by smoothly sliding it out. No hemorrhage was observed, and the jugular vein remained intact. The VAP and catheter were then removed (Fig 6). The subcutaneous tissue and skin of both incisions were closed with 3-0 PDS in simple, continuous patterns. Recovery from anaesthesia was uneventful, and no bleeding or bruising around the surgical sites was observed during the next 24 hours.
Interestingly, the bird's owners reported that, after discharge, the bird was able to stretch its neck more to crow, a behavior that had not been observed to the same extent while the VAP was in place.
This report describes the first known use of a VAP for the long-term administration of intravenous antimicrobial therapy in a bird. In this chicken, the VAP was used successfully with minimal complications throughout a 5-month period.
Pododermatitis is a common problem in poultry and raptors and occasionally in parrots and other species. Beginning as a superficial abrasion and ulceration of the plantar surface of the foot, it leads to cellulitis and abscessation in the deeper layers of the foot. Left untreated, it may progress to necrosis, tendonitis, septic arthritis, and osteomyelitis. Causative factors are body weight and condition, perch and flooring design, unequal weight-bearing because of unilateral lameness, and lack of exercise. Trauma to the plantar aspect of the foot then leads to secondary infections with pathogens including Staphylococcus aureus, Staphylococcus epidermidis, Corynebacterium species, E coli, Streptococcus faecalis, Pseudomonas species, Bacteroides species, Clostridium species, Candida albicans, and Aspergillus species. (1)
Several classification schemes exist for grading pododermatitis, but we prefer the 5-point system described by Bailey and Lloyd. (1) Using this system, this rooster's infection would be graded as a class IV pododermatitis (an infection of the deep, vital structures, producing tenosynovitis, arthritis, and/ or osteomyelitis, but retaining pedal function), carrying a guarded to poor prognosis for return to normal function.
Treatment of this class of pododermatitis involves debridement and appropriate antimicrobial therapy, as indicated by culture and susceptibility testing. However, any antimicrobial agent may select for resistance (2) and, in this case, the chicken had been previously been treated with various antimicrobial agents without an initial culture and susceptibility testing. Although these empirical therapies were not inappropriate, it seems likely that they may have inadvertently selected for a multidrug-resistant E coli.
Escherichia coli infections in chickens can usually be treated with potentiated sulfonamides, aminopenicillins, colistin, tetracyclines, spectinomycin, aminoglycosides, and enrofloxacin. (3) In Australia, fluoroquinolones and cephalosporins are not registered for use in poultry; in the United States, they are prohibited for use in food animals, including poultry, because of the creation of antimicrobial-resistant Campylobacter organisms. However, their use in backyard and pet chickens appears to be widespread around the world. Australia has adopted a conservative position regarding the registration of antimicrobials for food-producing animals and is the only country, to our knowledge, never to have permitted the use of fluoroquinolones. (4) Australian legislation also prohibits gentamicin use in production animals and places stringent label restrictions on the use of the third-generation cephalosporin, ceftiofur. Consequently, the level of antimicrobial resistance detected in chickens in Australia is low. (5) However, we are not aware of any studies that have examined the antimicrobial resistance profile of E coli from companion birds.
The E coli bacterium cultured was resistant to all commonly used drugs but was susceptible to amikacin, imipenem, and third-generation cephalosporins (ceftazidime, cefoxitin, and cefotaxime). These drugs are not absorbed after oral administration and, therefore, must be administered parentrally. (6-8) There are few studies with amikacin (9,10) and imipenem in chickens. Amikacin, an aminoglycoside, is potentially nephrotoxic and ototoxic, with the risk of toxicity increasing with the duration of therapy. (5) Imipenem, a carbapenem, is considered critically important for human health (11) and should only be used as a "last resort" drug in veterinary medicine. (4) Cephalosporins have few toxicity concerns, are bactericidal, and have good distribution to most tissues. (4) Ceftazidime and cefotaxime have also been previously recommended to treat infections from E coli in avian species. (12)
For these reasons, we decided to treat this chicken with ceftazidime, but this presented a practical problem. Although ceftazidime can be administered intramuscularly, the volume and frequency of injections and the likely duration of the treatment (2 mL of a 20 mg/mL solution q12h for several weeks or months) made iatrogenic muscle trauma and pain a likely consequence. For this reason, the use of a VAP was investigated.
Administration of long-term intravenous therapy in both people and animals is problematic. The use of intravenous catheters and injection ports is common but has the disadvantage in veterinary medicine of having most of the device on or above the surface of the skin, where self-removal of the device, occlusion, and phlebitis are likely consequences of long-term use. Consequently, their use is limited to days, rather than months, and frequent replacement is needed for ongoing treatments.
A VAP, or portocath, is a subcutaneously implanted device connected to a catheter, which is inserted into a central vein. The VAP consists of a chamber (the portal), covered by a silicone septum, which lies immediately under the skin and can be accessed by a noncoring Huber needle to allow the administration on intravenous drugs or the collection of blood. In medicine, VAPs are most commonly used for the administration of chemotherapy, for hematology patients, and for dialysis. Vascular access ports are becoming more frequently used in veterinary medicine, particularly for administering chemotherapy and for serial blood collection. (13,14) Senthilkumaran et al (15) reported the use of a VAP in chickens to obtain serial blood samples for hormone analysis. Of the 6 birds used in their trial, one developed a valvular endocarditis; the other 5 birds retained their VAPs for 3 months without incident, leading the researchers to conclude that VAPs are a safe and reliable method for maintaining intravenous access.
In people, the most common complications associated with VAPs are infection around the VAP, port migration, catheter occlusion, skin necrosis over the VAP, port exposure, device rotation, and catheter-related sepsis. Less common complications are catheter rupture, cardiac tamponade, catheter disconnection, and difficult catheter removal. (16)
In people, patient evaluation for systemic infection and preplacement antibiotic therapy are combined with strict asepsis in placing and maintaining the VAP to keep infection rates low, about 3%-7%. Skin necrosis and port exposure are more common in very thin patients, whereas port rotation is more common in obese patients. Care in selecting the placement site, ensuring the VAP is truly subcutaneous, and securely anchoring it in place will help to minimize these complications. (16)
Catheter occlusion can be due to kinking of the catheter or the formation of a clot within the lumen. Radiologic evaluation of the device after placement may be needed to evaluate the catheter for kinking, and clot formation can be minimized by regular flushing and then locking of the catheter with heparinized saline. The manufacturer (Norfolk Vet Products) provides directions with each device on the volume of heparinized saline to use for the various-sized ports. They recommend the VAP should be flushed with sterile saline and then locked with heparinized saline q24h for the first 3 days after surgical placement, then flushed and locked every time it is used. When not in use, a maintenance flush and lock every 3-4 weeks is usually sufficient to prevent occlusion. In this bird, the owners performed a maintenance flush and lock twice weekly for 3 months after treatment was discontinued; the catheter was still patent at the time of removal.
Although VAPs are more commonly used in mammalian patients, they potentially lend themselves to the treatment of avian patients requiring long-term intravenous therapy or serial blood collection. Vascular access ports offer the advantages of ease of access, reduced trauma and handling of the patient, and the accurate delivery of large volume or tissue-irritant drugs. The thin skin of birds may predispose the device to exposure and infection through self-mutilation, and this needs to be considered when selecting both the patient and the placement site. Provided they are implanted and maintained carefully, minimal complications should be expected.
(1.) Bailey T, Lloyd C. Raptors: disorders of the feet. In: Chitty J, Lierz M, eds. BSAVA Manual of Raptors, Pigeons and Passerine Birds. Hoboken, NJ: J Wiley and Sons; 2008:176-189.
(2.) Gyles CL. Antimicrobial resistance in selected bacteria from poultry. Anim Health Res Rev. 2008;9(2):149-158.
(3.) Lohren U, Ricci A, Cummings TS. Guidelines for antimicrobial use in poultry. In: Guardabassi L, Jensen LB. Kruse H, eds. Guide to Antimicrobial Use in Animals. Carlton, Australia: Blackwell Publishing Ltd; 2008:126-142.
(4.) Cheng AC, Turnidge J, Collignon P, et al. Control of fluoroquinolone resistance through successful regulation. Australia. Emerg Infect Dis. 2012;18(9):1453-1460.
(5.) Obeng AS, Rickard H, Ndi O, et al. Antibiotic resistance, phylogenetic grouping and virulence potential of Escherichia coli isolated from the faeces of intensively farmed and free range poultry. Vet Microbiol. 2012;154(3-4):305-315.
(6.) Prescott JF. Other [beta]-lactam antibiotics: [beta]-Lactamase inhibitors, carbapenems, and monobactams. In: Giguere S, Prescott JF, Dowling PM, eds. Antimicrobial Therapy in Veterinary Medicine. 5th ed. Ames, IA: Wiley-Blackwell; 2013:175-187.
(7.) Dowling PM. Aminoglycosides and aminocyclitols. In: Giguere S, Prescott JF, Dowling PM, eds. Antimicrobial Therapy in Veterinary Medicine. 5th ed. Ames, IA: Wiley-Blackwell; 2013:233-255.
(8.) Prescott JF. [beta]-Lactam antibiotics: cephalosporins. In: Giguere S, Prescott JF, Dowling PM, eds. Antimicrobial Therapy in Veterinary Medicine. 5th ed. Ames, IA: Wiley-Blackwell; 2013:153-173.
(9.) el-Gammal AA, Ravis WR, Krista LM, et al. A. Pharmacokinetics and intramuscular bioavailability of amikacin in chickens following single and multiple dosing. J Vet Pharmacol Ther. 1992;15(2):133-142.
(10.) Itoh N, Kikuchi N, Hiramune T. Antimicrobial effects of amikacin therapy on experimentally induced Salmonella Typhimurium infection in fowls. J Vet MedSci. 1996;58(5):425-429.
(11.) [WHO] World Health Organization. Critically important antimicrobials for human medicine. In: Report of the 3rd Meeting of the WHO Advisory Group on Integrated Surveillance of Antimicrobial Resistance (AGISAR), 2011. Oslo, Norway: WHO; 2012.
(12.) Fudge AM. Diagnosis and treatment of avian bacterial disease. Semin Avian Exot Pet Med. 2001;10(1):3-11.
(13.) Farrow HA, Rand JS, Burgess DM, et al. Jugular vascular access port implantation for frequent, long-term blood sampling in cats: methodology, assessment, and comparison with jugular catheters. Res Vet Sci. 2013;95(2):681-686.
(14.) Valentini F, Fassone F, Pozzebon A, et al. Use of totally implantable vascular access port with mini-invasive Seldinger technique in 12 dogs undergoing chemotherapy. Res Vet Sci. 2013;94(1):152-157.
(15.) Senthilkumaran C, Peterson S, Taylor M, Bedecarrats G. Use of a vascular access port for the measurement of pulsatile luteinizing hormone in old broiler breeders. Poult Sci. 2006;85(9):1632-1640.
(16.) Gonda SJ, Li R. Principles of subcutaneous port placement. Tech Vase Interv Radiol. 2011;14(4):198-203.
Robert James Tyson Doneley, BVSc FANZCVS, Bruce Austen Smith, BVSc, MSc, Dipl ACVS, FANZCVS, and Justine Susanah Gibson, BVSc, PhD
From the University of Queensland Veterinary Medical Centre (Doneley, Smith) and the School of Veterinary Science (Doneley, Smith, Gibson), The University of Queensland, Gatton, Queensland 4343, Australia.
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|Title Annotation:||Clinical Report|
|Author:||Doneley, Robert James Tyson; Smith, Bruce Austen; Gibson, Justine Susanah|
|Publication:||Journal of Avian Medicine and Surgery|
|Article Type:||Clinical report|
|Date:||Jun 1, 2015|
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