experimental animal urine collection

Collection of urine from experimentalanimals is a basic requirement in severalbiochemical, nutritional, urological,metabolic, toxicological, general behaviouraland physiological studies. The essentialfeatures of experimental animal urinecollection are: (1) obtaining pure urinewithout contamination with faeces oranimal feed, (2) collecting urine without anydirect intervention, (3) ease and convenienceof collection, (4) efficiency of collection and

(5) rapidity of collection (for simple spoturine tests).

Animal urine is collected either forpurposes of qualitative (commonly a clinicalveterinary purpose) or quantitative (mostlyan experimental scientific purpose) analysis.Urinalysis is one of the most informatoryand commonly performed laboratory testsavailable to practising veterinarians. Urinecollected for veterinary purposes is usefulfor ascertaining the general health andphysiological status of an animal, andhelpful in making an appropriate diagnosis.A small quantity of urine will suffice forqualitative urinalysis, which includes the

REVIEW ARTICLE

Experimental animal urine collection: a review

Biji T. Kurien1, Nancy E. Everds2 & R. Hal Scofield1,3,4

1Arthritis and Immunology Program, Oklahoma Medical Research Foundation, 3Department ofMedicine, University of Oklahoma Health Sciences Center, 4Department of Veterans Affairs MedicalCenter, Oklahoma City, Oklahoma, OK 73104 and 2Haskell Laboratory for Health and EnvironmentalSciences, Newark, DE 19714–0050, USA

Summary

Animal urine collection is a vital part of veterinary practice for ascertaining animal healthand in scientific investigations for assessing the results of experimental manipulations.Untainted animal urine collection is very challenging, especially with small rodents, and isan almost impossible task under conditions of microgravity. The fundamental aspects ofurine collection are: (1) ease of collection, (2) quality of sample, (3) prevention of contamina-tion, (4) severity of procedures used, (5) levels of pain caused to the animal and (6) refine-ment of methods to reduce stress, pain or distress. This review addresses the collection ofurine for qualitative and quantitative purposes from rodents, rabbits, felines, canines, avianspecies, equines, porcines, ungulates and certain non-human primates, with animal welfarein mind. Special emphasis has been given to rodents, canines and non-human primates,since they are the animals of choice for research purposes. Free catch (voluntary voiding),methods with mild intervention, surgical methods, modified restraint, cage and specialrequirement methods have been reviewed here. Efforts need to be taken to provideappropriate animal husbandry and to nurture the animals in as natural an environment aspossible since experimental results obtained from these research subjects are, to a greatextent, dependent upon their well-being. A continuous refinement in the procedures forcollecting urine from experimental animals will be the most efficient way of proceeding inobtaining pure urine specimens for obtaining reliable research data.

Keywords Animal urine; rodents; non-rodents; metabolism cages; animal welfare

Accepted 9 March 2004 © Laboratory Animals Ltd. Laboratory Animals (2004) 38, 333–361

Correspondence to: Biji T. Kurien, OMRF, MS # 24, 825 NE13th Street, Oklahoma City, OK 73104, USAE-mail: [email protected]

measurement of urinary pH, protein,glucose, bilirubin, haemoglobin, ketone,urobilinogen and creatinine levels.Quantitative excretion analysis for scientificpurposes requires the collection of timedurine samples (e.g. 24 h samples) and isuseful for ascertaining the result of anyspecial experimental manipulation that theanimal has been subjected to, when theurine collection has been standardized withrespect to time/water intake and all otherprocedural aspects. This is generally carriedout for the following purposes: (a) investiga-tion of renal function, (b) study of renaldiseases, (c) evaluation of metabolic and/orendocrine anomalies, (d) evaluation ofnutritional and metabolic requirements and(e) evaluation of excretion of a xenobioticand its metabolites.

We have undertaken this review to give abroad understanding of general proceduresavailable for collecting urine from variousanimal species and to critically review thebest method of collection for each species.The fundamental features of experimentalanimal urine collection are: (a) obtainingpure urine without contamination withfaeces or animal feed, (b) collecting urinewithout any compulsion and (c) ease andconvenience of collection. This review willaddress urine collection using the followingperformance criteria: (1) ease of collection,(2) degree of invasiveness or severity ofprocedures used, (3) quality of sample, (4)prevention of contamination and (5)different levels of pain, suffering or distresscaused to the animal. This review will alsoaddress the refinement of methods,especially the use of metabolism cages, toreduce stress, pain or distress. Refinementhas been defined as the minimizing ofanimal suffering during an experiment, andalso as the enhancement of animalwell-being through better husbandry(Morton 2002).

Many methods reviewed here would fallunder the category of ‘veterinary’ purpose,and have been sub-categorized under urinecollection without intervention/with mildintervention (the terms ‘free-catch’,‘voluntary voiding’ and ‘micturation’ havebeen used interchangeably for such collec-

tion). However several methods have beenincluded that use metabolism cages or otherspecialized equipment to collect urine forprescribed periods (such as 24 h periods)from which scientific data can be procuredto monitor specific interventions.

This review will address the collection ofurine for qualitative and quantitativepurposes from rodents (Table 1), rabbits,felines, canines, avian species, equines,porcines, ungulates and certain non-humanprimates (Table 2), with animal welfare inmind.

Collection of urine from rodents

Mice and rats are two of the primarymammalian species used for the evaluationof acute and chronic toxicity, metabolic, andbioavailability in preclinical evaluation ofdrugs and preregistration evaluation ofchemicals. They have proven useful onaccount of their defined genetic lineage, lowcost, relatively short lifespan and theminimal holding space required. Micecomprise a majority of all experimentalmammals among all animals used inresearch, teaching and testing. Mice areeasy to handle, inexpensive to maintain,genetically similar to humans with a shortlifespan and rapid reproductive rate, thusaccounting for the species position as theexperimental model of choice in research.Another primary use of mice is for theproduction of biological reagents, such asmonoclonal antibodies and vaccines. Miceand rats are used to test new procedures anddrugs for safety, as required by an array offederal regulations. The following descrip-tion includes some of the methods used tocollect urine from mice, rats and otherrodents (Table 1).

Urine collection without intervention(mouse)

Mouse urine collection using clear plasticwrap Kurien and Scofield (1999) describedtwo different methods for collecting puremouse urine for qualitative analysis. Thesingle animal method (Fig 1) involved

334 Kurien, Everds & Scofield

Laboratory Animals (2004) 38

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Animal urine collection 335




allowing a single mouse to urinate on Glad®

cling wrap, placed on white sheets of paper,outside the animal cage. The multipleanimal method (Fig 2) involved partitioningseven mice into seven different makeshiftcompartments (two X-shaped structuresfabricated with cardboard) laid out on topof the cling wrap and allowing them tourinate. The voided urine, in each case, was

then aspirated into microcentrifuge tubesusing an adjustable, air displacementpipette. Pure urine could be obtained in asquickly as 12 s without any compulsion.Volumes in the range of 10–250 �l wereobtained. Two rectangular pieces ofcardboard, each about 61 cm long and 22.9 cm wide, were used to make anX-shaped structure. To make this, a cut of5.1 cm long is made from the top middlesection of one of the cardboard pieces and acut of about 17.8 cm long from the bottommiddle of the other cardboard piece. Boththe cardboard pieces were linked to eachother in the middle at the cuts in such away that they fit snugly and gave theappearance of an X-shaped structure. TwoX-shaped structures gave an XX structure(Fig 2). The animals were placed within theconfines of these makeshift compartments.As soon as the animal urinated, the animalwas removed and the urine aspirated (Fig 2).

Urine collection with mild intervention(mouse and rat)

Mouse or rat micturation over Petri dishThis procedure involves holding the mouseor rat over a Petri plate or a plastic dispos-able weigh boat and encouraging it tomicturate (Watts 1971). Fresh, clean samplesfor the analysis of the entire urinary tractcan be obtained by expression of the bladdermanually into a cup or onto a tabletop byapplication of gentle trans-abdominalpressure over the bladder to overcomenormal urethral pressure. One micturationfrom a mouse was shown to produceapproximately 30 to 100 �l of urine, whichwas collected in a disposable plastic Petriplate, and housed in a humid chamber tominimize evaporation. Weigh boats arenon-wettable, and thus urine collected inweigh boats can be poured off into a testtube, which can be capped immediately.Generally, spontaneous urine collection isbest performed immediately upon enteringthe animal room.

Spot urine collection using a polystyrenebeaker (rat) Sometimes it becomes impor-tant to obtain urine from a large number of

Fig 1 Single animal method of mouse urine collec-tion using plastic wrap

Fig 2 Multiple animal method of mouse urine collection using plastic wrap

animals for qualitative analysis, e.g. todetect the presence or absence of glycosuria.For such purposes it becomes tedious andexpensive to use metabolism cages. Amethod was devised (Khosho et al. 1985) tocollect spot urine from rats that was simple,reliable and efficient.

A 5 ml polystyrene beaker (diSPo BeakerPolystyrene, American Scientific Products,McGaw Park, Illinois, USA) was attached tothe perineal wall using tape that wasadherent on both sides (Fig 3). This allowedthe collecting beaker to be attached andremoved with ease. The rat was then heldby the tail, after the beaker was attached, andthe back of the animal was stimulated tac-tilely using the fingers of the opposite hand.It was found that 80% of the animals voidedbetween 0.1 and 0.8 ml of urine within a fewseconds, while it took 5–10 min for theremaining 20%. Shaving and washing of theperiurethral area was found to be necessary ifthe collection of clean, debris-free urine wasessential. Hand holding of the beaker wasfound to be faster than attaching the beakerwith tape, but that procedure was found to besusceptible to urine spillage and contamina-tion with faeces.

Capillary tube method of urine collection(rats) Hayashi and Sakaguchi (1975)showed that capillary tubes could be used tocollect urine from male Sprague-Dawleyrats. The rat was held with one hand andthe lower part of the abdomen, around theurinary bladder, was pressed with the thumband the third finger of the other hand of thecollector to cause urinary excretion. Theurine excreted was immediately collecteddirectly into two capillary tubes heldbetween the index and middle fingers. Themethod was found to be suitable for routineurinalysis in rats and mice. However, it wasnot directly possible to estimate the volumeof urine collected using this procedure.

Modified restraint used for 24 h urinecollection

Polyethylene funnel method Perline (1971)designed an inexpensive apparatus (Fig 4) forcollecting mouse urine with the use of apolyethylene funnel (22.5 cm diameter). Anylon screen (14 mesh/in. cut in 12.5 cmdiameter) was inserted into the funnel con-caved and a stainless steel screen (4 mesh/2.5 cm cut 11.25 cm diameter) was placedover the upturned edges of the nylon screen.The screens were fastened to the funnel by



Fig 3 Illustration of a method to collect urine froma rat. The animal is tickled on the back and theurine is collected into a polystyrene beaker

Fig 4 A simple and inexpensive mouse urine collection device

Tab

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drilling six sets of two holes around theperimeter of the funnel and securing it bywire. The excess overlapping nylon screen-ing was trimmed. After placing a mouse inthe funnel, a plastic lid with ventilationholes punched into it was placed over thetop of the funnel to contain the mouse. Theapparatus was placed in an appropriate holderand a collection beaker placed beneath it.The cost of fabricating the apparatus wasestimated to be less than US$2.

Disposable individual mouse urinecollection apparatus Smith et al. (1981)constructed an inexpensive disposable appa-ratus, costing less than US$1, to collectindividual mouse urine using a disposable250 ml conical polypropylene centrifugetube and wire cloth. This procedure couldprove useful if highly toxic or carcinogenicagents are to be administered to the mice.Dilution of urine samples by the use of aleaky water bottle and contamination byfood have been eliminated by this apparatuson account of the use of external ports forwatering and feeding.

Essentially the conical end of the cylinderwas cut off to make a one centimetreopening in the cone bottom. A horizontal4 mesh (0.625 cm squares) galvanized wirecloth 5 �6 cm was positioned through slits inthe side wall of the cylindrical portion of thetube (9.5 cm from the top), upon which themouse was allowed to rest during the periodof urine collection. Another wire grid (corrosion-resistant chrome steel wire cloth:16 mesh, 1/16 square) was inserted throughslits in the lower portion of the tube to retainthe faeces while allowing the urine to flowthrough the conical portion of the tube. Twoopenings, covered with 4 mesh galvanizedwire cloth, were made to provide access tofood and water. The excreted urine flowedinto a tube held in place in a container filledwith ice so that the urine was maintained at4°C. Urine volumes ranged from 0.5 to 2.0 mlper 24 h per 25 g mouse.

An apparatus for 24 h mouse urinecollection Jones et al. (1973) constructedan apparatus to collect individual mouseurine. It is similar to that of Perline (1971)

but possesses certain advantages like allow-ing easier cleaning, having reduced surfacetension, and allowing greater freedom ofmovement for the mouse with a lower levelof stress. It comprised a Gridweld(17.5 �17.5 �15 cm) animal cage 20 cm2

24-gauge aluminum funnel with a 0.94 cmdiameter hole at the apex and a polyethyl-ene tube (12.5 cm length and 0.78 cm indiameter). The animal cage rests inside thefunnel, and a 6 mesh/2.5 cm stainless steelgrid is placed beneath the cage inside thefunnel. The funnel and the grid are dippedin silicon to reduce surface tension andfacilitate the passage of urine. The poly-ethylene tubing rests in the collection vesseland protrudes into the funnel thus prevent-ing the passage of faecal pellets into thecollection vessel but without hampering the passage of urine. The plasticine preventsmovement of the collection vessel.

The mouse is placed in the apparatus andallowed to acclimatize for 24 h before urinecollection was started. A routine the authorshave found successful, involved urinecollection for 16 h followed by feeding for8 h. The funnel, etc., were removed duringthe feeding period to eliminate contamina-tion with food particles (Table 1).

Commercial metabolism cage methods for24 h urine collectionNumerous commercial metabolism cagesare available to contain experimentalanimals for the purpose of collecting urine(Harvard Apparatus, MA, USA; BraintreeScientific, MA, USA; Techniplast, NJ, USA).These cages are meticulously engineered toeffectively separate faeces and urine intotubes outside the cage. The separation isimmediate and complete with no urinewashover and no chance for urine to enterthe faeces tube, yielding untainted andreliable samples. A typical commercialmetabolism cage is constructed as follows.A chamber made of transparent, gnaw-proofpolycarbonate with removable vented covershouses the animal. It has a feeder chamberlocated outside the cage that contains afeeder drawer that slides out for easy fillingwith slurries, liquids or powders, without



disturbing the animal. This set-up preventsthe urine from getting contaminated withthe feed. The small size of the externalchamber discourages the animal fromnesting or sleeping inside. The cages have awater bottle support and spillage collectiontube designed to prevent water fromentering the cage and contaminating theurine. The spillage collection tube iscalibrated and thus allows an investigator tomeasure the actual water intake by theanimal. The faeces and urine collectiontubes are made of non-wettingpolymethylpentene and allow the faecalpellet or the urine to flow down intoseparate collection tubes. The cages areconstructed in such a way that the urinenever washes over into the faeces tube. Eachexcreta can be removed without causing anydisturbance to the animal. This allows teststo be conducted over a prolonged periodwithout interruptions caused by animalre-acclimatization. In addition, all cagecomponents are autoclavable. The metabo-lism cage requires a single-cage stand, whichcould be used with either the small rodentor large rodent cages.

In spite of the fact that different kinds ofmetabolism cages have been described for thecollection of urine from experimental ani-mals, it has not been clearly shown that thecollection procedures are not stressful for theanimals. This is particularly true in animalsthat are placed in cages with floor areas thatare much smaller than those recommendedfor the keeping of experimental laboratoryanimals (Merkenschlager & Wilk 1979). Also,the metabolism cages are constructed fromwire-mesh bases, which could meanuncomfortable conditions for the animal oreven tissue injury to feet or legs, probablyresulting from pressure sores (Ewbank 1984,Lawlor 2002).

The importance of animal welfare forobtaining good experimental results hasbeen stressed recently as a result of severalscientific studies (Anderson et al. 1972,Hughes & Black 1976, Miller et al. 1986,Jezierski & Konecka 1996, Lensink et al.2000, Rochlitz 2000, Russell 2002) and bythe European Economic Council Directivein the 1980s (European Economic

Community 1986) and more recently by the American Association for LaboratoryAnimal Science (AALAS 2001). ‘Stressedanimals do not make good research subjects’has been stressed time and again (AmericanMedical Association 1992, cited byReinhardt and Reinhardt 2000).

Historically, it has been the norm tohouse rodents in typically small, barren andmonotonous laboratory housing (highlystandardized conditions) to reducevariability in data and responses in researchinvestigations. Recent evidence suggeststhat housing them in such conditions affectsthe animal fundamentally so that even theresults obtained from studies using suchhousing are being questioned (Wurbel 2001,Sherwin 2002). The emphasis in animalhousing has been on changing to a‘performance’ approach (providing housingconditions that help the animals to attaincertain performance standards) from an‘engineering’ approach (specifying cagedimensions and features) (National ResearchCouncil 1996). Other studies have shownthat the welfare of rats could be improvedby using solid cage flooring instead of gridcage flooring (Manser et al. 1995, Manseret al. 1996). Studies have also shown thata more complex housing environment,compared to the barren cage, buffers anxietyresponses to potential stressors (Levine1985). Rats with access to an appropriateshelter have been found to be more explo-rative and less timid than those in barrencages (Townsend 1997). Manser et al. (1998)have found that nest boxes of opaque orsemi-opaque materials are particularlysuitable shelters. The current system ofhousing does not cater to the behaviouralrequirements of the animal, other than thebasics of feeding and drinking. Laboratorymice have been shown to show a diversebehavioural repertoire (Jennings et al. 1998),which is thwarted when they are housedunder conventional laboratory housing.Consequently, these mice frequently exhibitso-called abnormal behaviours (e.g. stereo-typies) indicating that mice do experiencechronic frustration when placed in conven-tional caging. Moreover, the sensorycapabilities of mice have been rarely



considered in housing and husbandry design,which has been compared to rearing ofanimals under conditions of sensorydeprivation or interference (Sherwin 2002).

Alternatives to commercial cages forscientific purpose

Single-mouse urine collection and pHmonitoring system Commerciallyavailable mouse metabolism cages havelarge collection areas that result in poormouse urine recovery efficiencies andcontamination by faeces and feed. Westet al. (1978) designed a glass cage withminimal surface area to house mice for24 h urine collection, for use in conjunctionwith a flow-through pH electrode and lowtemperature bath to monitor urinary pH andcollect samples.

The apparatus consisted of a cylindricalglass cage 13 cm high and 5.8 cm indiameter, with a funnel-shaped bottomfitted with two stainless steel screens (top:8 mesh wire cloth, 5.7 cm diameter; bottom:12 mesh, 5.7 cm diameter) to support theanimal and prevent the faeces from lodgingon the urine collecting surface (Fig 5). Thescreens were separated by a stainless steelhoop one centimetre high and 5 cm indiameter. Three holes were bored into theglass walls of the cage. One hole allowedaccess to food. The second hole supplied awater sipper tube, while the third hole,positioned by the upper screen, was used by

the mice to discard faeces adhering to theupper screen. A cup-shaped cover 8 cm indiameter and 3 cm high with 0.8 cm holeswas used for ventilation purposes.

An experiment performed with a radiola-belled compound excreted in the urinerevealed a recovery efficiency of 74.2%,while 25.8% of radioactive material wasfound adhering to the cage surfaces. A flow-through pH electrode, attached to the cagewith a 1.5 cm piece of 2 mm interiordiameter latex tubing, used in combinationwith a digital pH meter and strip chartrecorder, provided a continuous pH versustime urination record.

Aluminium foil method Black andClaxton (1979) devised a method to collecturine from rats caged in hanging-type cagesusing aluminium foil attached to its bottom,approximately 10 cm from the front of thecage and hanging 1.5 to 2.5 cm below thefloor of the cage (Fig 6). Wrinkles in the foilprovided small spaces for the urine to accu-mulate, keeping it free from the faeces.Collection of urine samples (approximatelyone millilitre) was found to be virtuallycomplete within 4 h. This method was notsuitable for overnight collection, as theinvestigators found that the rats attemptedto chew the foil, if they could make contactwith it. In addition, the urine was found toevaporate, and faecal material accumulatedsuch that it was difficult to obtain unconta-minated urine.



Fig 5 Urine collection cage for a single mouseFig 6 Hanging type rat cage with aluminium foilat the bottom for collection of urine samples

Fenske (1989) devised a way to collect24 h urine from small experimental animalslike Mongolian gerbils, rats, guineapigs andtree shrews by keeping them in cagesidentical in size to their home cages duringthe collection period. Fenske placedaluminium sheets below cages equippedwith stainless steel mesh floors andcollected 24 h urine quantitatively fromthese sheets. In spite of the fact that urinedried up in patches, it was found that thedried urine could be effectively removedfrom the aluminium sheets by a singlewashing step (recovery rate: 84–100%).

One hundred per cent separation methodJackson and Sutherland (1984) fabricated anew urine collecting device for quantitativelycollecting urine that (a) provided a 100%separation of faeces from urine, (b) gave aquantitative collection of small urinesamples (0–7 ml), (c) was devoid of metal inits design and (d) was resistant to organicsolvents. The apparatus has a mountingplate (made from either acetal resin or acetalresin coated with polymethyl methacrylate)with a projecting funnel to receive the penisof the rat (or urethral region for femaleanimals), the plate being adhered to thepelvic skin by a quick-drying methylcyano-acrylate adhesive. The funnel and urinecollector are separate units joined by screwthreads disposed around a cylindrical projec-tion or annulus at the mouth of the funnel.Screw threads couple with screw threads inan opening through the top wall of thecontainer, with the container being essen-tially a chamber that is vented in the topwall. A drain positioned in the end wall isused to empty the container. Urine couldbe sampled with a syringe or by merelyremoving the container from the funnel.

Urine collection for pharmacokinetic anddrug metabolic studies Toon and Rowland(1981) fabricated a device out of Plexiglas torestrain chronically cannulated rats andcollect faeces-free urine during chronicpharmacokinetic and drug metabolicstudies. The main part of the device is aclear Plexiglas tube (35 cm long with aninternal and external diameter of 6.3 and

7.6 cm, respectively) with a slit (0.4 cmwide) cut into the top to accommodateexteriorized cannulae, which enablesampling and drug administration. Analuminium diamond mesh (4.5 �35, meshsize 0.3 cm) positioned at a height of onecentimetre from the bottom of the tubeformed the floor of the tube. The floor isremovable, for purposes of cleaning, and isheld in place during use by three stainlesssteel pegs. These pegs protrude from thebottom of the tube and fit into holes boredinto three Plexiglas strips (4 �0.6 �0.6 cm)which are themselves secured by screws tothe mesh floor. One Plexiglas strip is posi-tioned 2 cm from each end of the floor witha third strip positioned under the middle ofthe floor. To prevent loss of urine, the openends of the tube beneath the mesh floorare blocked using a one centimetre deepPlexiglas segment. A stainless steel tube(one centimetre diameter, 3 cm long) whichacts as a drain to permit urine collection, ispositioned through the base of the tube atone end.

Special requirements for quantitativedetermination of unstable compounds

Low-temperature urine collection Urinespecimens have to be frozen soon after void-ing for the quantitative determination ofunstable compounds present in urine so thatoxidative, enzymatic and bacterial degrada-tion can be minimized. Denckla (1966,1969) developed a method to collect freshly-voided rat urine by free fall through a wiremesh flooring onto a �55°C refrigeratedplate or into dry ice-cooled heptane. In theunit that he developed in 1969, the urineand faeces fall onto heptane cooled dry ice(Fig 7). The main container is a picnic coolerhaving a linear polyethylene sheath over aninner core of expanded styrene foam. Anadditional 5.1 cm of fibreglass insulation isused on the sides and bottom of the unit.The urine-collecting pan is a Teflon®-coatedbaking pan of suitable size to fit freelyinside the cooler. The pan has stringsattached to facilitate removal and handling.A stainless-steel mesh (approximately0.078 cm) strainer is riveted to one to



drain out the heptane, and a 2.5 cm hole isdrilled in the bottom of the pan for aneoprene stopper for the collection of urine.The rat cage is supported approximately3.75 cm above the edge of the cooler on asimple aluminium frame with notchedwooden feet resting on the cooler’s top edge.About 4 kg of dry ice was found to last fora 16 h run. As the dry ice sublimes, the pansinks slowly and therefore always containsheptane approximately 2.5 cm thick. Urine

forms spherical pellets and these do notadhere to faeces. The faecal pellets can beeasily removed with forceps before the urineis collected.

Lartigue et al. (1978) fabricated a morecomplicated modular low-temperature urinecollection apparatus for the collection ofrodent urine at �19°C (Fig 8). The unitconsisted of a covered and insulated metalpan in which propylene glycol was cooled bya standard cooling probe and circulated by alow-speed stirrer. The pan (70 �70 �9 cm)was constructed of stainless steel. A metalsleeve was welded into the centre, near thebottom, of one wall of the pan. This accom-modated a length of insulating tubing(Armaflex, Armstrong Cork Co, Lancaster,PA) designed to fit snugly around the flexi-ble hose of a cooling probe (Cryocool, CC-60F, Neslab Instruments, Portsmouth, NH).The inner sleeve-tubing junction wassecured by silicone rubber adhesive, and theexternal tubing-flexible hose junction by acircular hose clamp. The pan was filled withpolyethylene glycol to completely immersethe cooling probe. For insulation, theexterior of the pan was covered with sheetsof polystyrene 2.5 cm thick. The rim of thepan protruded one centimetre above the8 cm high insulation. A 75 �75 cm sheet ofpolystyrene grooved on the underside to a



Fig 7 Low temperature apparatus for collectingurine. A pan containing heptane and cooled by dryice is used to collect the urine

Fig 8 Cross-section of the low temperature mouse urine collection apparatus showing the details of con-struction

depth of one centimetre at a distance of2.5 cm from the four edges formed a snuglyfitting cover. Holes, 21 cm in diameter, werecut in the cover to accommodate five rodentmetabolism cages. The holes were spacedin such a way that they did not interferewith the cooling probe. An oblique hole wasdrilled near the cooling probe to accommo-date a stirring rod. The stirring rod’s motorwas mounted 35 cm above the lid on a hori-zontal support. This distance was requiredfor the cage units to be changed. To improvecirculation of polyethylene glycol under thecage units, three spacers 0.5 cm in heightwere placed on the pan floor beneath eachhole. The cage units were inserted using awater–food deflector ring to cover the smallspace between the side of each cage and thehole. A hole was drilled through the topnear one side of the pan to accommodate athermometer.

A plastic bulb suspended from the lowerend of the funnel deflected faeces to thebottom of the cage and allowed urine toflow over the surface of the bulb into acollection cup at the bottom. This methodallowed the collection of urine volumesup to 50 ml per collection period and provided good separation of faeces andurine. However, the large mesh flooring ofthe cage did not totally prevent coprophagy(stool eating). Tail cups were used (Frape et al. 1970) if total faecal collection or the elimination of coprophagy is a requirement.

Collection of mouse or rat urine in themicrogravity of space Gravity holds us tothe ground, keeps the moon in orbit aroundthe earth, and the earth in orbit around thesun. It is a force that governs motionthroughout the universe. It is commonnotion, however, that there is no gravityabove the earth’s atmosphere, i.e. in ‘space’,and consequently there appears to be nogravity aboard orbiting spacecrafts.Normally, orbital altitudes involving humanspaceflight vary between 120–360 miles (192to 576 km) above the surface of the earth. Inthese regions, the gravitational field is stillquite strong, since this is only about 1.8%of the distance to the moon. The gravita-

tional field of the earth at about 250 miles(400 km) above the surface maintains 88.8%of its strength at the surface. Therefore, thespace shuttle or space station is kept inorbit around the earth by gravity.

Sir Isaac Newton first described thenature of gravity more than 300 years ago. Itis the attraction between any two masses,which is most apparent when one mass isvery large (like the earth). Normal gravity,or 1 g, is the acceleration of an object towardthe ground caused by gravity alone, near thesurface of the earth, which is equal to32.2 ft/s2 (9.8 m/s2).

If an apple is dropped on Earth, it falls at1 g. The apple that an astronaut drops on thespace station falls too, but it just does notlook like it is falling. This is because theyare all falling together—the apple, theastronaut, and the station. However, thedifference is that they are not fallingtowards the earth but falling around it.However, since they are all falling at thesame rate, objects inside of the stationappear to float in a state which is calledzero gravity (0 g), or more accurately micro-gravity (1 �10�6g) (National Aeronauticsand Space Administration, USA).

Several space flight models, to be used inearth-based research, are available to mimicthe physiological changes seen in the ratduring weightlessness as seen under condi-tions of microgravity (Musacchia et al. 1980,Wronski & Morey-Holton 1987). However,information on nutrition or gastrointestinaland renal function in the space flight modelreported by Wronski and Morey-Holton, themost widely used animal model, has beenlimited by the difficulty of obtaining uncon-taminated urine specimens for analysis(Harper et al. 1994). In the Holton system, atraction tape harness is applied to the tail,and the rat’s hindquarters are elevated byattaching the harness to a pulley system.Weight-bearing hindlimbs are unloaded, andthere is a headward fluid shift. The tail-suspended rats are able to move freely abouttheir cages on their forelimbs and toleratethis procedure with a minimum of stress.In this system, however, urine, faeces andspilled food fall through the grid floor ontoabsorbent paper beneath the cage and cannot



be separated and recovered quantitativelyfor analysis in metabolic balance studies.

Harper et al. (1994) have described amodified system where they have addeda separator system to collect urine and faecalsamples for metabolic and nutritionalstudies in the tail suspension model. Thebasic design for the cage top and suspensionapparatus remained the same as that of theHolton model. However, in this newermodel, the food cup was relocated from aninside corner of the cage floor to extendoutside the cage front through a 3.8 cmlong �5.1 cm outer diameter tube with agrid floor. The animal can reach throughthe tunnel for food, and any spilled foodfalls through holes in the plastic grid of thetunnel floor onto a small tray set beneaththe floor. This food can be returned to thefood cup prior to weighing to determine foodconsumption. A funnel at the base of thecage has a tab on the exterior of the funnelspout, and to this the urine and faeces sepa-rator is attached. The separator collectsurine and faeces into a 30 mm diameterscrew-top centrifuge tube with a 30 or 50 mlcapacity by means of a linear diffuser. Theurine flows down the funnel and onto thelinear diffuser where narrow channels leadthe urine to the opening of the collectiontube. The faeces roll down the funnel andover these channels into a second tube.Complete engineering specifications andsuppliers for materials are given in a NASAtechnical memorandum (Evans et al. 1994).This apparatus has been successively used tocollect post-flight urine after the SpacelabLife Sciences-2 space shuttle flight.

Water droplets can float about under theconditions of microgravity in space, creatinga potential hazard for electrical equipment.The space shuttle system has been designedto incorporate air pressure and flow (toreplace gravity) for the pneumatic collectionand transport of only human wastes intowaste processing equipment. Mouse or raturine collection is not currently practicalin the microgravity of space, and Kurienand Scofield (1999) have suggested a hypo-thetical way to obtain small volumes ofurine by placing the animals in tetheredziplock bags (or similar transparent bags)



fitted with tubes to deliver oxygen and toaspirate voided urine.

Surgical methods

Urine collection from anaesthetized malerats Urine collection from rats hasgenerally been either through the use ofmetabolism cages (Sunderman 1944, Harnedet al. 1949, Peacock & Harris 1950) or bysurgical cystostomy. White (1971) developeda method for collecting urine from anaes-thetized rats utilizing an external drainagecatheter. After anaesthesia with subcuta-neous urethane, a 200–400 g male rat isplaced supine and restrained. Digitalcompression alongside the base of theforeskin forces the rat’s penis to protrude.A modified polyethylene Luer-Endintramedic catheter is then fitted over thetip of the penis. Then, a haemostat is usedto pull the foreskin over the penis catheterjunction for fixation with a silk ligature.The catheter is then withdrawn fromagainst the tip of the penis to where thesecuring foreskin ligature is just distal tothe catheter’s end. This prevents urethralobstruction, which results if the catheter isfixed tight against the penis. When fluidsare forced via an orogastric tube, a urineflow of 6–10 cc per hour is obtained. Urinecollected using this method was found to befree from cells and comparable to normallyvoided urine.

Urine collection from other smallanimals

Exotic pet medicine is a field that ischanging continuously. A number ofdifferent species are being marketed eachyear and these animals originate fromdifferent corners of the world. Veterinarycare becomes very important in these pets,since the pet owners are largely ignorantregarding the basic husbandry, nutritionalneeds and medical needs of many of theseanimals. Urine is an important diagnosticsample for the determination of renal func-tion and urinary tract infection (Ness 1999)in these animals.

Ness (1999) mentions different ways inwhich urine can be collected from exoticsmall animals, such as the Chinchilla(a rodent native to the cool dry climate ofthe Andes), prairie dog (a rodent approxi-mately the size and shape of a guineapig),the hedgehog and the sugar glider (a small,arboreal, nocturnal marsupial). Urine can besiphoned off from the cage floor with easeprovided the sample is free from contamina-tion with faecal material, which is criticalfor the correct interpretation of results.Urethral catheterization is not generallypractical for collecting urine in smallanimals and in such cases cystocentesis(described later on) could be used.

Urine collection from rabbits

Urine collection with mild interventionGarvey and Aalseth (1971) showed thaturine samples could be easily collected fromnewborn rabbits (12 h to 10-day-old rabbits).The rabbit is cradled on its back with itshead toward the wrist in the left hand of theoperator, whose thumb presses down on theleft hind foot. The rabbit is not allowed tostruggle by firmly gripping the rabbit. Withthe thumb of the right hand the abdomen ofthe rabbit is stroked gently but with slightpressure. The strokes should begin at thestomach level and end just beyond thebladder area. The most pressure should beexerted in the region of the bladder area.Stroking should be maintained until urine isexpelled, which begins only when themuscles are relaxed. After 10 days, restraintof the rabbit must be adjusted. Theoperator’s thumb and little finger grasp theanimal around the neck and rib cage, andthe hind feet are hand free. Stroking of theanimal is as mentioned earlier. However, alonger period of stroking is necessary beforeurine is released as a result of increasedmuscular control. The bladder function isno longer stimulated at this juncture andtherefore it is necessary to isolate the rabbitfor several hours to permit bladder fillingprior to collection. The experimenter has toremember that too much pressure orrestraint could lead to discomfort, internal



injury or suffocation. In addition, individualrabbits vary in the time required for relax-ation and therefore several minutes ofstroking may be necessary. Up to 5 ml ofurine has been collected using this method.

Urine collection by cystocentesisCystocentesis is a procedure by whichsterile urine can be obtained directly fromthe bladder, without potential contamina-tion from the distal urethra or genital tract,by inserting a small needle. Cystocentesis inrabbits is carried out as in cats (described inthe next section). Essentially, the bladder islocated in the caudoventral abdomen, whichcan be felt when full. A sterile syringe ispassed through the abdominal wall into thebladder to collect the urine sample directlyin the bladder. This is the ‘cleanest’ way toprocure a urine sample, as it is obtaineddirectly from the bladder and will be free ofbacteria endemic to the urethra (Table 2).

Urine collection using metabolism cagesRabbit urine collection for quantitativepurposes has been achieved by usingcommercial metabolism cages (24 h urinecollection) (Benson & Paul-Murphy 1999).Typically a rabbit cage should provide forsocial or environmental enrichment suchthat it would provide them an avenue toindulge in some normal behaviour such assitting up on the hind legs, hopping, diggingand hiding. In the past, laboratory rabbitshave been usually kept in small cages,which have been shown to have detrimentaleffects (Boers et al. 2002). Provision ofadequate space (to allow for normal posturaladjustments with freedom of movement),straw bedding, wooden sticks and treebranches (for gnawing), cardboard boxes andplastic crates or sections (to serve as substi-tute burrows and ‘safe’ places to retreat)have been found to be very important. Someof the hay should be placed on top of thecage to enable the animal to spend someextra time retrieving it through the bars ofthe cage. Provision must be made so that anindividual animal is not visually separatedfrom other rabbits (since rabbits love to live

in groups). The primary enclosure of single-housed animals should be at least 75 cm by80 cm in dimension. The base of the cagesshould not have grid or wire flooring,since that would result in ulcerativepododermatitis. The urine collection fromthe rabbits should be as stress free aspossible.

Urine collection from felines

The best way to obtain sterile urine from acat is by cystocentesis. The second bestcollection method is ‘free-catch’, where theurine is collected in a sterile or cleancontainer as the urine is leaving the body.The third, but least desirable, method is thecollecting of the urine out of a container oroff the floor after it has been voided.Obtaining of urine by free catch or by thethird method is less stressful and painfulto the animal than cystocentesis. For qua-litative purposes, like measurement of pHand other metabolites like glucose, protein,etc., it may suffice to collect urine by the‘free catch’ method.

CystocentesisCystocentesis can be performed on smallerspecies with the use of a 25 gauge or smallerneedle. Additionally, cystocentesis can beperformed on the exposed bladder afteranaesthesia immediately prior to necropsy.

The importance of cystocentesis, both asa diagnostic and therapeutic tool, has beenrecognized for over 80 years. Diagnosticcystocentesis circumvents many of thepotential problems associated with thecollection of urine specimens by normalmicturation, manual compression of theurinary bladder, or catheterization (Krugeret al. 1996) in feline patients with non-obtrusive lower urinary tract diseases(LUTD) or those with non-urinary disorders.

Cystocentesis can be performed in severalways. It can be done with the animal lyingdown on its back with its limbs facingupwards (dorsal recumbency), in lateralrecumbancy, standing or while being heldstanding on its hindlimbs by elevating its



forequarters. Cystocentesis is usuallyperformed with a 25–22 gauge needle. Thehair at the puncture site is clipped and thearea is cleaned with an antiseptic solution.The bladder is first palpated and immobi-lized. Care should be taken to ensure thatthe bladder is not squeezed tightly whilethe puncture is being made since this cancause urine to leak from the puncture siteinto the abdominal cavity. The needle isinserted at a 45° angle, a short distancecranial to the junction of the bladder andurethra. The needle should be inserted intothe bladder while the plunger is pulled backto create a negative pressure. However, ifurine is not obtained, the needle should notbe reinserted, to avoid the risk of penetrat-ing a bowel loop and subsequently insertingthe contaminated needle into the bladder.The needle has to be changed before its rein-sertion into the bladder. If urine is notobtained after three attempts it may be dueto the fact that the bladder is probably smalland situated in the pelvic canal. Uponobtaining the urine sample, the syringeplunger is released, and the needle removedfrom the abdomen (Dhein 2002).

Other than inducing mild transient micro-scopic haematuria (which may be indistin-guishable from the pathological haematuriaassociated with many naturally occurringfeline LUTDs) cystocentesis has not beenassociated with significant side effects.

Cage methods for quantitative purposes

Cage collection When clean, daily caturine samples were required and when facedwith the non-availability of metabolismcages, Matandos and Franz (1980) deviseda method to collect urine from caged labora-tory cats. They used this method to collecturine from 34 cats housed individually infibreglass cages equipped with perforated,stainless steel floors elevated 8 cm above thefibreglass floor. A small board (3 �2 �35 cm)was attached to the bottom of a modifiedround rubber dish pan (37 cm diameter �13 cm deep) such that the pan tilted 6°from the horizontal (for drainage). A hole(8 mm diameter) was punched into the panat the lowest point in the outer circumference

of the base of the pan. The barrel of a 3 mldisposable syringe was cut transversely atthe 0.25 ml mark and pushed through thehole in the pan from the inside. To assurecomplete drainage of the pan, the fingergrips on the plunger end of the syringe barrel were notched with a small hand-heldelectric grinder. The syringe barrel wascemented in place with epoxy liquid adhe-sive after a good fit was achieved. For betteradhesion, the surfaces of both the pan andthe syringe were roughened with fine sand-paper before the application of the epoxy.The modified pan was placed in the backcorner of the cage with the drain hole awayfrom the corner. The syringe-barrel drain,protruding through the floor, held a 100 mldisposable beaker in place under theelevated cage floor. The authors have usedthis technique for more than 200 cat-daysand had a collection success rate of morethan 82%. The cat urinating outside the panor not urinating at all accounted for 9% ofthe failures, while loose faeces contami-nating the urine, faeces blocking the drainhole or the pan being moved from over thebeaker accounted for 3% failure in eachcase.

Solid floor caging Cats are normallyhoused in relatively small metabolism cageswith wire-mesh bases, which makes it veryuncomfortable for the animals, especially ifthey are to be confined in them for longperiods. These cages with wire-mesh floorsprevent the cats from burying their faeces.Confined cats have been shown to preferburying their faeces (Bateson & Turner1988). Suppression of such behaviour couldinterfere with the cat’s welfare, thereforePastoor et al. (1990) have developed an alter-native method for collecting urine from catsusing metabolism cages with solid floors,equipped with modified litter boxes to sepa-rate urine and faeces and allowing cats tobury their faeces.

Cats are more likely to respond to poorhousing conditions by inhibiting normalbehaviours (feeding, grooming, exploring andplaying) or by becoming inactive since theydo not have as wide a behavioural repertoire(facial expression, body posture) for visual



communication as the highly social, group-living dog (Rochlitz 2002). Keeping cats,therefore, in a species-appropriate environ-ment will make it easier for urine collection(or for other scientific investigations) inaddition to enhancing animal welfare (Poole1997). The primary enclosure used for catsmust have enough space for them to expressa wide range of normal postures andbehaviours, such as stretching, exploringand playing, and to allow the investigatorto carry out routine procedures, like urinecollection, easily. It is the quality of thecage space rather than the quantity that isimportant for the cats, once certainminimum size criteria are fulfilled (Rochlitz2000) (Table 2).

Urine collection from ferrets

Cystocentesis is the method of choice forcollecting urine from these animals, whenthe urinary bladder is palpable (Morrisey &Ramer 1999). The urinary bladder can beaccessed through the lateral body wall, as ina cat, with a 3 ml syringe and 2.5 cm gaugeneedle. Proper restraint is vital for thisprocedure and distracting the animal with asweet substance has been found to besuccessful. Urethral catheterization hasbeen found to be difficult in these animals.

Urine collection from canines

Specimens of dog urine for bacteriologicalculture and urinalysis have been collectedusing three methods, namely voluntaryvoiding, cystocentesis and catheterization.

Voluntary voiding using mild interventionUsing digital compression of the bladder,Carter et al. (1978) collected voluntarilyvoided samples, as clean as possible, insterilized cups. Occasionally, however, theyfound that the cups made contact with hairin the area of the vulva of female dogs or theprepuce of male dogs. The entire voidedurine could be collected only occasionally,and in a majority of cases the samples werecollected midstream. Studies have shown,

however, that samples collected midstreamcould be contaminated with bacteria origi-nating either from the urethra or adjacent tothe external urethral orifice. In one studyinvolving 32 dogs (sex distribution notgiven), urine samples collected from 25 dogshad bacterial growth (Klausner et al. 1975,Finco & Kern 1977, Carter et al. 1978). Thisresults in the misdiagnosis of urinary tractinfection (UTI). In a second study, 15 of 25dogs with no evidence of UTI had bacterialcontamination in the urine collected bymidstream catch (Finco & Kern 1977). In yetanother study involving 50 clinically normaldogs (25 males and 25 females), 85% ofvoluntarily voided urine specimens hadbacterial contamination (Comer & Ling1981). Urine from female dogs gave a higherpercentage of bacterial contamination (91%).

CystocentesisCystocentesis has been used (Carter et al.1978) to collect urine from dogs. First thedogs were sedated with acepromazine(Ayerst Laboratories, New York, NY, USA).The hair in an area of the abdominal wallbetween the midline and the flank wasclipped and the area was cleansed cleanwith alcohol, soap and mild tincture ofiodine. The bladder was first immobilizedusing digital palpation, and then a 5-mlurine sample was drawn by using a syringefitted with a 22 gauge, 3.25 cm needle. Atthe time of ovariohysterectomy the sampleswere aspirated from the bladder followingexposure by laparotomy.

Urine specimens obtained from dogs usingcystocentesis were found to be bacteriologi-cally sterile (Klausner et al. 1975, Finco &Kern 1977, Carter et al. 1978). In one studyinvolving 50 clinically normal dogs (25males and 25 females), urine specimensobtained from all the dogs using cystocentesiswere found to be bacteriologically sterile.

CatheterizationVarious groups have used catheterization tocollect urine from dogs (Carter et al. 1978,Mulcahy et al. 1978, Russel et al. 1987).Catheterization, using sterilized plastic



cathethers lubricated with sterile lubricatingjelly (Lubafax, Burroughs Wellcome Co.Research Triangle Park, NC, USA), has beenused to collect urine samples from dogs.The end of the penis in the males or theperivulvar area in the females was wipedwith gauze sponges saturated with organiciodine solution and wiped dry. The initial5 ml of urine withdrawn through thecatheter, in males as well as females, wasdiscarded and a new syringe was attached towithdraw samples for purposes of culturingfor bacteria.

Mulcahy et al. (1978) implanted adurable cystostomy cannula in the urinarybladders to collect urine for prolonged peri-od in awake dogs. The implantation of the cannula was performed by simplecystostomy in female dogs. This was notperformed in male dogs as the penis waslocated at the site of the cystostomy. Thedogs were found to initially pick at thecannula with their teeth and paws.However, since the plugs of the cannula arecovered with caps fastened with stainlesssteel screws, they soon learn that they areimpossible to remove. All urine gravitatesreadily to the cannula, with the animal inan erect position, and exits immediatelyfrom the bladder. Irrigating the bladder toremove all urine is therefore unnecessaryand this becomes very useful when accurateurine volumes for osmolality determina-tions must be obtained.

A procedure for enabling multiplecatheterizations using a bowel window hasbeen described (Finkle et al. 1961) for dogsthat have undergone ureterosigmoidostomy.This is a procedure that implants the ureterinto the sigmoid colon, resulting in thediversion of urine into the bowel. This iscarried out as a consequence of bladderdisease or cystocentesis. Urine collectionfrom these animals poses a serious problemsince the urine will be contaminated withfaeces. To circumvent this problem, ‘abowel window of Lucite’ (DuPont) wasdevised (Finkle et al. 1961). The bowelwindow was surgically installed into thesigmoid colon. After further surgical proce-dures a No. 4 French woven-silk ureteralcatheter was directed up the ureter until

clear urine made its appearance. Minor leak-ages occurred transiently around the window,and low-grade infection was found to occur.These catheterizations sometimes initiatedpyelonephritis. In spite of these problems,these investigators have found that manyof their experimental animals survived morethan 100 days postoperatively, followingnumerous ureteral catheterizations.

Prolonged indwelling urinary cathetershave been shown to result in UTIs in dogsand cats (Barsanti et al. 1985) and risk ofinfection was shown to be increased withduration of catheterization. This catheter-associated infection occurred despite antibi-otic therapy.

Catheterization as such is not an asepticprocedure in spite of the fact that theurethral orifice is cleansed thoroughly priorto insertion of the catheter. The urethras ofmale and female dogs have resident popu-lations of bacteria, many of which arepotentially pathogenic. Cross-contaminationwith these bacteria would lead to mislead-ing results. Also, catheterization of the blad-der in people has been associated with UTI.In separate studies it has been found thaturine samples obtained using catheteriza-tion were found to be positive for bacterialculture (Klausner et al. 1975, Finco & Kern1977, Carter et al. 1978, Biertuempfel et al.1981, Comer & Ling 1981). Catheterizationof the bladder, in humans, has been shownto cause UTI resulting from the contamina-tion and subsequent colonization of the blad-der mucosa with bacteria originating fromthe urethra (Biertuempfel et al. 1981).

The dog has been associated with man fora very long time and is one of the oldestdomesticated animal. It has a special statusamong people and is often considered as amember of the family. Therefore, when adog is used as an experimental subject itreceives special protection in several coun-tries. In the United Kingdom, for example,special justification has to be providedbefore dogs can be involved in a studyinvolving pain, suffering or distress. Dogsare inquisitive animals and will react badlyto barren or sensorily restricted environ-ments. It has been well established thathousing environments that do not cater to



the physical or social needs of an animallead to alterations in physiology andbehaviour, which ultimately influences theresearch data. Therefore a dog kennel designshould (a) keep the animal in good health—physically and mentally, (b) allow easyhandling of the dog by the personnel, (c) belarge enough to allow group housing ofcompatible dogs, (d) be flexible to allowpens to be combined together to makelarger runs, (e) allow choice of location and provide interest within the enclosure,(f ) provide some shelter from kennel matesthrough the use of visual barriers and(g) minimize sound egress (Hubrecht 2002).

Avian urine collection

Avian urine collection is useful for identifi-cation of microorganisms like parasites,bacteria, fungi or viruses. It also hasdiagnostic merit for the identification ofdrug metabolites, sex steroids, pathologicalprocesses (haematuria, haematochezia) andtoxins (Greiner & Ritchie 1994, Arnold &Holt 1996, Hagedorn et al. 1997).

Urine collection in the fowl presents apeculiar problem since the urine flows fromthe ureters into the cloaca, where it mixeswith the faeces. A variety of methods havebeen used to separate urine and faeces. Oncea fresh dropping is obtained on a cleannon-porous surface, urine can be carefullyseparated from the faeces. The capillaryaction of a plain microhaematocrit tube oraspiration via a needle and syringe has beenused to collect the urine, leaving faecesbehind (Murray 1997, Styles & Phalen1998).

Non-surgical means of collecting urine byinserting a funnel into the urodeum havebeen employed (Davis 1927, Coulson &Hughes 1930) and the cannulation of theureters has been employed by Hester et al.(1940). The catheter (funnel) and cannulatechniques facilitate only the collection ofurine for limited periods of time. Hart andEssex (1942) and then Ainsworth (1965)have described the exteriorization of theureteral openings, which require several

incisions and sutures. It also requires that abird wears a harness to facilitate urineand/or faeces collection. Hart and Essex(1942) described construction of an externalanus, which remained suitable only for 2 or3 weeks.

The quantity and character of the urineexcreted by birds have usually been vital towork on other problems concerned withavian renal physiology. The studies on thesebirds have not been very systematic. Hesteret al. (1940) have studied urine secretion inthe chicken in order to get a thoroughunderstanding of the daily output volumeand the character of the urine secreted bybirds, under a variety of conditions. Theycarried out three series of experiments:(a) the ureters were cannulated under localanaesthesia and the effect of certaindiuretics and other drugs were observed;(b) urine, free of faecal material, wascollected by inserting a small funnel havingsufficient diameter to include the twoureteral orifices into the urodeum and(c) collection of urine under near normalcondition as possible was made possible byexteriorization of the ureteral orifices by anoperation.

The following are two classic surgicalmethods used to collect urine from fowl.

(a) Separation of urine and faeces in theimmature fowlColvin et al. (1966) described a surgicalmethod that circumvented many disadvan-tages of previous methods. White-Leghorncockerels (9 weeks of age) were fasted for8 h to insure that all faeces were voidedfrom the lower portion of the gut. Afteranaesthesia, feathers were removed fromthe vicinity of the cloacal orifice. Anincision was made perpendicular to themidline and about one centimetre anteriorto the anal orifice. The distal portion of therectum was brought to the surface of theincision and the cloaca was closed bymeans of a silk suture as near the rectalsphincter as possible. The rectum was thensevered just proximal to the suture. At thispoint, a few milligrams of powderedsulfathiazole was dusted into the bodycavity through the incision. A sterile,



pliable, rubber medicine-dropper bulb witha rolled end was cut to a length of approxi-mately 2.5 cm and the cut end was inserted1.5 cm into the rectum. The tube was thensutured to the rectum by six sutures nearthe end of the inserted end. The mucosawas then attached to the surface of the skinimmediately around the incision by silksutures and dusted with sulfathiazole.A tube as described above was inserted intothe cloaca about 0.75 cm so as not to blockthe ureteral openings. The tube was thenattached to the dorsal and ventral lips ofthe cloaca by six silk sutures.

In order to collect urine or faecal samples,toy balloons of a suitable size were tied withnylon string around the rolled end of therubber bulbs which protruded from thecloacal orifice and the artificial anus. Birdstreated surgically in this way were found torequire virtually no postoperative attentionand could move around freely in about 1–2 hand be given food and water immediately.The authors found that it was necessary towait about 3 days before placing the birds onexperimental treatments in order to ensurethat the diuresis observed by Hester et al.(1940) and Hart and Essex (1942) would beovercome and to ensure that the sutureswould hold the tubes securely in place. Inabout 10% of the birds, recurring blockageof the tubes was observed by the authors,who discarded such birds. By using extrudedtubes the collection balloons could beattached and removed easily without theinconvenience of a harness on the bird, andcontamination of the faeces by the urine isprevented.

(b) Collection of urine from the chickenAs mentioned earlier, collection of uncont-aminated avian urine specimens is difficultbecause the urine of the bird flows fromthe ureter into the cloaca where it can mixfreely with faecal matter. Several earlierinvestigators have devised a variety ofmethods for obtaining avian urine free ofdejecta. Minowski (1886) avoided faecalcontamination of geese urine by ligatingthe rectum above the cloaca. Milroy (1903)suggests that this does not work well, andhe in turn attempted to obtain uncontami-

nated urine by using a specially devisedtube designed to divert the faeces from theportion of the apparatus that was to carryoff the urine. Since faecal contaminationcould not be prevented this way he thenseparated the rectum and cloaca by anoperation. He made an artificial anus in theanterior abdominal wall and urine wascollected into a suitable bag attached belowthe anal orifice for as much as 24 h (volumeranged from 500 to 1500 ml during thisperiod). Paton (1910) made an artificialanus just like the one Milroy made butinstead of collecting in a bag, he collectedit in a metabolism cage. Paton states in hiswork that the amount of 24 h urine couldnot be accurately estimated since the birdsspilled some of the drinking water into it.Sharpe (1912) anaesthetized hens underurethane or ether and collected urine bycannulating the ureters. Mayrs (1924) alsoobtained urine from cockerels by cannulat-ing the ureters under urethane anaesthesiaand obtained an urine output of 0.69 ml/min from an average of seven experiments.Davis (1927) collected urine by means of acatheter inserted into the urodeum suchthat the urine was drained off as it flowedfrom the ureteral orifices. He inserted cotton plugs in the rectum, sometimes, to prevent faecal contamination. He foundthat he could obtain more urine underanaesthesia. He found that without anaesthesia, the urine was so concentratedthat the solids precipitated and clogged the catheter.

Urine collection from miniature pigs

Miniature strains of pigs, as well as farmstrains, have been used extensively inlaboratories. Small pot-bellied pigs have alsobecome common pets in both urban andrural households resulting in an increasingneed for providing health care to theseanimals.

As pets, the miniature pigs are largelyhousebroken and therefore they have to betaken outdoors to obtain urine sample by vol-untary voiding (Van Metre & Angelos 1999).Since these animals possess remarkable speed



and agility, they can be difficult to catchonce they escape, and therefore placingthem in a harness and a leash is vital. Urinesamples from females are easier to obtaincompared to males. The pendulous abdomenof these pigs makes it difficult to collecturine from males. Urine has been collectedfrom these animals using the cap from a60 ml syringe case fastened to an unwoundcoat hanger to serve as a urine collectiondevice. The cap was moved under the pig’sabdomen once urination began to obtain aurine sample.

It has been difficult to catheterize thebladder of the male miniature pigs owing tothe presence of the urethral recess (this is asmall hollow of the urethral lumen locatedon the dorsal aspect of the pelvic segment ofthe urethra, at the level of the ischium). Asthe catheter is advanced through the urethrait tends to become lodged at the recess. Ithas been possible for the veterinarian,however, to direct the catheter into thebladder by placing a finger into the pig’srectum and gently pushing the catheterventrally and cranially (Van Metre &Angelos 1999).

Pigs have to be kept well and carefullyhandled, on account of the fact that theyare conspicuously sensitive animals, toobtain reliable research data. While housingpigs, care must be taken to provide themwith roughage such as oat hulls and strawso that they can perform foraging activitiesand fill their gut. A barren concrete floormay result in foot and joint irritations andinflammation. Therefore the animals mustbe maintained on straw bedding. Pigs arevery social in nature and therefore theymust be maintained in compatible pairs andalso should be exposed to positive humancontact in their home pens on a regularbasis. Even playing a radio with a variety oftalk and music has been shown to be help-ful for the pigs to get accustomed tohumans. When a pig is isolated for obtain-ing a sample or used for a research proce-dure, another familiar pig needs to be takenalong with it. This will prevent them fromgetting apprehensive, agitated or distressedwhen separated from their herdmates(Grandin 2002).

Urine collection in large animals

Metabolic stalls for restraining male cattle,sheep, goats and swine and collecting urineand faeces separately are available. Stalls areavailable that use gratings, screens and fun-nels to separate faeces and urine and theseare satisfactory for many purposes and easyto use. However, in these stalls, cross-contamination of excreta occurs in varyingdegrees. Other systems, such as indwellingcatheters and apparatus attached to animalsby harness arrangements, have been devel-oped to minimize or prevent cross contami-nation of excreta. However, these methodssometimes cause animal discomfort orbladder infection, or the apparatus does notstay in place for extended urine collection(Aschbacher 1970).

Urine collection from male cattle, sheep,goats and swinePaulson and Cottrell (1984) constructeda simple apparatus to collect urine fromthese animals that circumvented theseproblems. The apparatus was constructed ofelastic, waterproof cloth, fabric fastener anda waterproof bag. It was lightweight, easilyinstalled on the animal and resulted in noapparent discomfort to the animal and wasfound to be suitable for extended collectionperiods.

Urine collection from female ungulatesFor determining physiological responses ofanimals to changes in a variety of environ-mental stimuli such as ambient tempera-ture, energy and nutrient levels and wateravailability, metabolic studies with rumi-nants are important. Such studies generallyrequire the measurement of feed offered, andthe separation and quantification of urineand faeces produced by individual animals.Total collections of urine and faeces aredifficult to achieve from female cattle andbison since the proximity of the meatusurinates and ishio-rectal region causescross-contamination of samples. As a result,metabolic studies often use male subjectsbecause of the obvious ease of separatingand collecting urine and faeces. If total urine



collections from female ruminants arerequired, bladder catheterization is thetechnique most commonly used. However,catheters present unique problems such astheir difficulty of insertion, secondarybladder infections and often substantialurine loss. Deliberto and Urness (1995)developed a urine collection apparatusmodelled after the urine deflector flapdeveloped for faecal collection bags byKartchner and Rittenhouse (1979). Thismethod was also designed to collect urineunlike the urine deflector flap (Table 2).

Collection of urine from equines

Collection of urine in the horse is limitedto voluntary voiding and catheterization.Cystocentesis is not possible practicallyowing to the size of the horse and also dueto the intrapelvic position of the equinebladder.

Equine urine collection often forms anintegral part of studies on water balance(Groenendyk et al. 1988) and renal function(Rawlings & Bisgard 1975, Lane & Merritt1983, Kohn & Strasser 1986) in manyresearch establishments. Investigations onwater homeostasis in equines are usuallycarried out in metabolic stalls equipped withfloor pans to simplify the collection of free-flow urine samples. Variations of an appara-tus, to collect urine from male horses,suspended beneath the penis by means of aharness have been described (Van der Nootet al. 1965, Tasker 1966, Harris 1988). Onlyone of these (Harris 1988) describes a non-invasive collection apparatus that could beused in mares. Urine from female horses hasbeen collected using indwelling bladdercatheters in studies involving renal clearanceof electrolytes (Rawlings & Bisgard 1975,Lane & Meritt 1983, Gronwall & Price 1985,Glade 1986, Kohn & Strasser 1986). Theseinvestigations involved introducing a sterileFoley catheter (28 or 32 French gauge) intothe bladder aseptically and keeping it inplace by inflating the balloon with 30 ml ofair or water. Urine was drained either con-tinuously from the bladder into a collectingreservoir or intermittently every 2 h through

the catheter (Kohn & Strasser 1986). Thismethod has the drawback of bacterialinfection, with one study showing three outof 13 horses having bacteruria at the end ofthe 12 h trial period (Rawlings & Bisgard1975).

Another method (Van den Berg 1996) forcollecting urine from mares used a devicemade out of rubber in the form of a skirt.The device was shaped in the form of achute by means of pliable metal on theoutside of the rubber. Using leather straps,the apparatus was attached to a backharness such that the proximal tip of thechute touched the mid-vulva region. Thefree end of the chute was then led to acollection funnel, between the hind legs,suspended in front of the hind legs. Thisdevice was used to collect urine routinely,safely and easily from female horses.

Collection of urine from non-humanprimates

Lesser, or grey, mouse lemurs (Microcebusmurinus), nocturnal owl monkeys (Aotustrivirgatus), common marmosets (Callithrixjacchus), squirrel monkeys (Saimirisciureus), capuchin monkeys (Cebusapella), owl monkeys (Aotus trivirgatus),rhesus macaques (Macaca mulatta),cynomolgus macaques (long-tailed macaques)(Macaca fascicularis), vervet monkeys(Ceropithecus aethiops), baboons (Papiospp.), bonnet macaques (Macaca radiata),pigtailed macaques (Macaca nemestrina),rhesus macaques (Macaca mulatta), chim-panzees (Pan troglodytes) and baboons aresome of the non-human primates used inresearch (Reinhardt & Reinhardt 2000,European Commission 2002).

New drugs are evaluated for preclinicalsafety in a rodent and a non-rodent (‘sec-ond’) species before controlled clinical trialsusing human patients or volunteers.Non-human primates are commonly used inthe safety evaluation of putative pharmaceu-ticals where other non-rodent species (e.g.dogs) have been demonstrated to be a lesspredictive model of the human responseowing to dissimilar kinetic, metabolic or



pharmacological responses, idiosyncraticreactions or lack of biological response. Thecommon marmoset (Callithrix jacchus) hasalso been used in preclinical safety evalua-tion studies for over a decade. Rats havebeen used as the rodent species in mostcases and the dog or non-human primates[macaque monkeys—cynomolgus (Macacafascicularis) or rhesus (Macaca mulatta)usually] are used for preclinical toxicologystudies.

Many primatological investigators arefearful of non-human primates and thereforecondone the forceful restraint of theseanimals during procedures that required anyform of physical human–non-humanprimate interaction. Squeeze backs are usedto immobilize an animal by pushing it tothe front of the cage, during blood or urinecollection. In addition, restrain chutes withadjustable restrictors and/or guillotinepanels are used to restrain an animal duringsampling procedures. Such involuntaryrestraint procedures introduce stress-relatedphysiological variables that can confoundresearch data. Being isolated from familiarhomecage surroundings during mostrestraint conditions adds to this distress.The British Code of Practice for the Housingand Care of Animals Used in ScientificProcedures (1989) points out the fact thattraining the non-human primate tocooperate during the procedure used is theleast distressing method of handling theseanimals.

Urine collection from marmosets andtamarinsHearn et al. (1975) reported a system for therestraint and collection of blood and urinefrom these animals. In their method, themarmoset was transferred to a rat metabo-lism cage for a 24 h period. Urine wasseparated automatically from faecal materialin the cage funnel and passed through a sidearm into a disposable plastic containerimmersed in ice.

The administration of radioactive-labelledmaterials for metabolic studies could not beundertaken using the above method owingto the fact that the cage volume was too

small to allow the animal to be held formore than 24 h. Metabolic studies stipulatedthe holding of the animal in the metabolismcage until the label was cleared primarilyinto the urine and faeces. Lunn (1989)modified the method of Hearn et al. for thispurpose. He modified a Metabowl III model10 glass metabolism cage (Jencons ScientificLtd, Leighton Buzzard, Bedfordshire, UK) toincrease its height, by the addition of astainless steel cylinder. In addition, heincluded a stainless steel feeding box insteadof the standard cigar type feeder and angledthe drop lick water bowl arm towards thecage interior to improve its accessibility.Animals were held in the Metabowl cage forup to 5 days continuously without anyapparent ill effects.

In another method, groups of marmosetswere familiarized with a custom-made urinecollection apparatus as an extension of theirhutch box (Anzenberger & Gossweiler1993), and were slowly conditioned to urinate right after the lights were turned on for a food reward. Experimenters foundthat, by the end of 3 weeks of training, theycould collect individual urine samples froman entire family of 4–16 animals.

Group-housed female tamarins (Saguinussp.) were conditioned without using aformal training protocol or investing extratime to urinate into containers shortly afterwaking up in the morning in return for afood reward (Snowdon et al. 1985).

Urine collection from the macaque monkeyLopez-Anaya et al. (1990) have used asimple method to collect urine fromnon-restrained infant macaques (Macacanemestrina). They have adapted a diapermethod, used in paediatric medicine, for thetotal urine collection for use in phamacoki-netic studies. A diaper/nappy was madeusing cellulose sponges, polyethylene sheetsand VelcroTM. First, a single layer of thesponge was cut to the shape of a diaper. Theregion to be positioned over the genitals wasconstructed with two sewn-together layersof sponge to maximize urine-absorptioncapacity. The sponge unit was positioned ina slightly larger, but similarly shaped,

polyethylene envelope such that the singlelayer part of the sponge was secured betweenthe polyethylene sheets by tape to preventcross-contamination of urine and faeces. Inaddition, an opening was made for the tail.VelcroTM fittings along the edges of the longextensions of the diaper kept the diaper inposition on the infant macaques. By weigh-ing the diaper before and after urination, theauthors were able to ascertain the volume ofthe urine obtained (assuming the specificgravity of urine to be unity). They squeezedthe diaper to obtain aliquots of urine for fur-ther assays. The authors have successfullyused this procedure to measure recovery ofdrug metabolites efficiently. The spongecould hold 20 times its weight in water,which allowed for urine collection over along period of time. In some cases, a smallamount of faecal material was found to bepresent in the sponges. They were not, how-ever, found to mix significantly with theurine, judging from the stability of one ofthe metabolites studied.

A Japanese group (Hayakawa & Takenaka1999) have collected urine fromindividually-caged Japanese macaques(Macaca fuscata) by placing a vat coveredwith a screen under their cages. Urinesamples were collected from the vat using adisposable plastic pipette. Urine volumesranging from 0.02 to 14 ml were obtained,with the smallest volume being from aninfant macaque.

Urine collection from the vervet monkeysand chimpanzeesTraining non-human primates to cooperateduring experimental procedures, using foodas a source of positive reinforcement, hasbeen found to be one of the most usefulenvironmental enhancement options.Group-housed male vervet monkeys(Ceropithecus aethiops) were trained byKelley and Bramblett (1981) to urinate ondemand, using peanuts as a reward, into abeaker. During this procedure, the animalswere not separated from their social group.Six of eight males were found to predictablyproduce urine samples after a 2-monthperiod of training.



Stone et al. (1996) trained 19 femalegroup-housed chimpanzees to cooperatewith urine sample collection. They foundthat the mean duration of training time toreach reliable performance was 237 minutes.

The housing system most commonly usedfor housing non-human primates is summa-rized as follows (European Commission2002):

(a) Individual housing in a single cage withfollowing specifications The cagesshould reduce risks of transmissionof diseases between animals and shouldbe amenable to easy cleaning andsanitation.

(b) Pair and connected cages for twoprimates or a small group of animalsThis caging system should permanentlyhouse compatible pairs of non-humanprimates and provide for the animal tospend part of the day in physical contact,by joining adjacent cages. This systemallows for the temporary separation ofanimals when sample collection orexperimental procedures are performed.

(c) Pens for group housing These are largecages, rooms or corrals enclosed byfences, walls, bars or meshed wire andare used generally for housing groups ofanimals. In such cases, for samplingprocedures, special capture procedures asoutlined by Reinhardt and Reinhardt(2000) may be required.

(d) Indoor–outdoor enclosures Here anoutside area is connected to an indoorheated space, where animals can findprotection from adverse weather andseek refuge from threats and attacks.

Comfortable housing for the non-humanprimates is just as important as in the caseof the other animals, to promote their well-being and therefore the quality of researchconducted with them. Social companionshiphas been found to be a necessity, just likefood. Housing the animals in pairs, startingat the juvenile stage, has been shown tobe very effective. The companion servesas a stress buffer in fear-provoking situa-tions like experiments that require chairrestraints or tether restraints. When theanimals have to be placed alone temporarily,



as during a metabolic study, the cagingarrangement must be such that it allows theindividual animal to maintain visual con-tact with at least one conspecific to reducethe stress of social deprivation (Reinhardt2002).

In addition, the physical as well as apsychologically comfortable environmentprovided to an animal plays an importantpart in obtaining reliable research findings.A trustful relationship with the attendingpersonnel coupled with compassionate carehas been shown to be very critical. Themacho-type person has been shown to trig-ger distress reactions in the animal thatresult in skewed experimental results(Reinhardt 2002). It has been aptly said that‘the behaviour of an animal during a proce-dure depends on the confidence it has in itshandler’ and that, ‘this confidence is devel-oped through regular human contact and,once established, should be preserved’(Home Office 1989). This sentiment isechoed by the National Research Council(1998). Many primates possess the capabilityfor learning to interact with a caretaker in acooperative manner, making it possible toteach them simple tasks such as voluntarilypresenting themselves for urine or bloodcollection. This kind of positive reinforce-ment training enables non-human primatesto become ‘partners’ rather than ‘victims’ inresearch.

Conclusion

Pure and reliable urinary samples are verychallenging to obtain from experimentalanimals, especially from small rodents. Inparticular, untainted animal urine collectionis almost an impossible task under condi-tions of microgravity. The numerousmethods for experimental animal urinecollection reviewed here should help readersto better understand the intricacies andvagaries involved in obtaining pure,uncontaminated urine.

Free catch (voluntary voiding), surgical,modified restraint, cage and special require-ment methods are some of the methods thathave been described for urine collection inrodents. Times and volumes of collection

varied with the different methods. Of all themethods we have reviewed we found onlyone method that did not use any kind ofintervention to collect urine from rodents(Kurien & Scofield 1999). Urine could becollected very rapidly and without anycontaminating species or feed, by using thismethod. Ingenious methods for the collec-tion of urine from larger animals, includingcystocentesis and catheterization, have beendescribed. All these methods have beeninstrumental in obtaining pure animal urinefor qualitative and quantitative studies.

Most of the methods that have beendescribed here, however, do not takeadequate precautions to provide appropriateanimal husbandry. Since experimentalresults obtained from these researchsubjects are, to a great extent, dependentupon their well being, it is of the utmostimportance to nurture them in as natural an environment as possible. In addition,compassion and good care should be showntowards all animals, be they rodents,canines or non-human primates.

Care must be taken to minimize pain anddistress to the animals as much as possible.For example, cystocentesis should be usedonly as a last resort when sterile urine isabsolutely essential. For qualitative studiesthat can be carried out rapidly, free catchshould be the method of choice. In general,from an experimental viewpoint it shouldbe regarded as standard practice to use the‘free catch’ method for collecting urinefrom rodents, rabbit, canines, minipigs andnon-human primates. A combination ofcatheterization and/or a metabolism cageshould be used for larger animals (e.g.domestic livestock).

There are relatively few laboratory specieswhere urine collections cannot be performedusing a metabolism cage. Some modifica-tions may be required in the general designto accommodate species-specific require-ments, but the key consideration is toprevent contamination of the urine samplewith food debris, water and/or faeces. Cagescan be readily modified to accommodatecannulated animals or even those that areexposed to a test atmosphere or whereexpired air is being collected. There are



relatively few situations where urine cannotbe collected.

A continuous refinement in the proce-dures for collecting urine from experimentalanimals will be the efficient way to proceedin obtaining pure urine specimens for gener-ating reliable research data.

Acknowledgments We thank Ms Marilyn Bonham-Leyba and Ms Beverly Hurt of the GraphicsResources Center, OMRF for their excellent effort indrawing the figures and formatting the tables pre-sented in this manuscript.

This work is supported by NIH grant ARO1844 toRHS and the Oklahama Center for the Advancementof Science and Technology to RHS and BTK.

References

American Association for Laboratory AnimalScience (2001) Cost of Sharing: RecognizingHuman Emotions in the Care of LaboratoryAnimals. Memphis, TN: American Association forLaboratory Animal Science

American Medical Association (1992) Use ofAnimals in Biomedical Research—The Challengeand Response. An American Medical AssociationWhite Paper. AMA. Group on Science andTechnology, Chicago, IL

Anzenberger G, Gossweiler H (1993) How to obtainindividual urine samples from undisturbedmarmoset families. American Journal ofPrimatology 31, 223–30

Ainsworth L (1965) Surgical procedure for exterior-ization of the ureteral openings of the hen. PoultryScience 44, 1561

Anderson CO, Denenberg VH, Zarrow MX (1972)Effects of handling and social isolation upon therabbit’s behaviour. Behaviour 43, 165–75

Arnold JW, Holt P (1996) Cytotoxicity in chickenalimentary secretions as measured by a derivativeof the tumor necrosis factor assay. Poultry Science75, 329–34

Aschbacher PW (1970) An adjustable metabolic stallfor cattle. Journal of Animal Science 31, 741–4

Barsanti JA, Blue J, Edmunds J (1985) Urinary tractinfection due to indwelling bladder catheters indogs and cats. Journal of the American VeterinaryMedical Association 187, 384–8

Bateson P, Turner DC (1988) Questions about cats.In: The Domestic Cat: The Biology of its Behavior(Turner DC, Bateson P, eds). Cambridge:Cambridge University Press, pp 193–201

Benson KG, Paul-Murphy J (1999) Clinical pathologyof the domestic rabbit. Acquisition and interpreta-tion of samples. The Veterinary Clinics of NorthAmerica. Exotic Animal Practice 2, 539–51, v.Review

Biertuempfel PH, Ling GV, Ling GA (1981) Urinarytract infection resulting from catheterization inhealthy adult dogs. Journal of the AmericanVeterinary Medical Association 178, 989–91

Black WD, Claxton MJ (1979) A simple, reliable andinexpensive method for the collection of rat urine.Laboratory Animal Science 29, 253–4

Boers K, Gray G, Love J, Mahmutovic Z,McCormick S, Turcotte N, Zhang Y (2002)Comfortable quarters for rabbits in research insti-tutions. In: Comfortable Quarters for LaboratoryAnimals, 9th edn (Reinhardt V, Reinhardt A, eds).Washington DC: Animal Welfare Institute

British Codes of Practice for the Housing and Care ofAnimals used in Scientific Procedures (1989)Home Office Animals (Scientific Procedures) Act1986. London

Carter JM, Klausner JS, Osborne CA, Bates FY (1978)Comparison of collection techniques for quantita-tive urine culture in dogs. Journal of the AmericanVeterinary Medical Association 173, 296–8

Colvin LB, Creger CR, Couch JR, Ferguson TM,Ansari MN (1966) A simplified method for separa-tion of urine and faeces in the immature fowl.Proceedings of the Society for ExperimentalBiology and Medicine 123, 415–17

Comer KM, Ling GV (1981) Results of urinalysis andbacterial culture of canine urine obtained byantepubic cystocentesis, catheterization, and themidstream voided methods. Journal of the AmericanVeterinary Medical Association 179, 891–5

Coulson EJ, Hughes JS (1930) Collection and analysisof chicken urine. Poultry Science 10, 53

Davis RE (1927) The nitrogenous constituents of henurine. Journal of Biological Chemistry 74, 509–13

Deliberto TJ, Urness PJ (1995) A total urine collec-tion apparatus for female bison and cattle. Journalof Range Management 48, 92–3

Denckla WD (1966) Low temperature urine collectorfor unstable compounds. Journal of Laboratoryand Clinical Medicine 68, 173–6

Denckla WD (1969) Inexpensive low-temperatureurine collector. Journal of Applied Physiology26, 393–4

Dhein CR (2002) Small Animal Diagnostic andTherapeutic Techniques. Cystocentesis.http://www.vetmed.wsu.edu/courses_samDX/cysto.htm

European Commission (2002) The Welfare of Non-Human Primates used in Research. Report of theScientific Committee on Animal Health andAnimal Welfare

European Economic Community (1986) Councildirective 86/609 on the Approximation of Laws,Regulations, and Administrative ProvisionsRegarding the Protection of Animals Used forExperimental and Other Scientific Purposes,Annex II Guidelines for Accomodation and Careof Animals. Official Journal of the EuropeanCommunities L358, 7–28



Evans J, Mulenburg GM, Harper JS, Skundberg TL,Navidi M, Arnaud SB (1994) A Metabolic Cage forthe Hindlimb Suspended Rat. NASA AmesResearch Center, pp 1–46

Ewbank R (1984) Laboratory dogs and cats: Report ofthe working group. In: Standards in LaboratoryAnimal Management. Potters Bar: The UniversitiesFederation for Animal Welfare, pp 105–6

Fenske M (1989) Application of a new, simplemethod for quantitative collection of 24-hoururines in small laboratory animals: determinationof basal excretion of proteins, creatinine, urea,electrolytes, and of free steroids. Zeitschrift furVersuchstierkunde 32, 65–70

Finco DR, Kern A (1977) Pyelonephritis. In: CurrentVeterinary Therapy VI (Kirk RW, ed).Philadelphia: WB Saunders, pp 1106–11

Finkle AL, Karg SJ, Smith DR (1961) Bowel-windowcollection of urine after ureterosigmoidostomy indogs. Journal of Applied Physiology 16, 752–5

Frape DL, Wilkinson J, Chubb LG (1970) A simpli-fied metabolism cage and tail cup for young rats.Laboratory Animals 4, 67–73

Garvey JS, Aalseth BL (1971) Urine collection fromnewborn rabbits. Laboratory Animal Science 21, 739

Glade MJ (1986) Estimation of urinary flow rate inweanling and yearling horses. American Journalof Veterinary Research 47, 2151–4

Grandin T (2002) Comfortable quarters for pigs inresearch institutions. In: Comfortable Quartersfor Laboratory Animals, 9th edn (Reinhardt V,Reinhardt A, eds). Washington DC: AnimalWelfare Institute

Greiner EC, Ritchie BW (1994) Parasites. In: AvianMedicine: Principles and Application (Ritchie BW,Harrison GJ, Harrison LR, eds). Lake Worth, FL:Wingers Publishing, pp 1007–29

Groenendyk S, English PB, Abetz I (1988) Externalbalance of water and electrolytes in the horse.Equine Veterinary Journal 20, 189–93

Gronwall R, Price G (1985) Estimation of urine flowrate in mares. American Journal of VeterinaryResearch 46, 1107–10

Hagedorn HW, Zankl H, Grund C, Schulz R (1997)Excretion of the anabolic steroid boldenone by rac-ing pigeons. American Journal of VeterinaryResearch 58, 224–7

Harned BK, Cunningham RW, Gill ER (1949) Ametabolism cage for small animals. Science109, 489–90

Harper JS, Mulenburg GM, Evans J, Navidi M,Wolinsky I, Arnaud SB (1994) Metabolism cagesfor a space flight model in the rat. LaboratoryAnimal Science 44, 645–7

Harris P (1988) Collection of urine. EquineVeterinary Journal 20, 86–8

Hart WM, Essex HE (1942) Water metabolic of thechicken (Gallus domesticus) with special refer-ence to the role of the cloaca. American Journal ofPhysiology 136, 657–68



Hayashi S, Sakaguchi T (1975) Capillary tube analysisof small animals. Laboratory Animal Science25, 782–3

Hayakawa S, Takenaka O (1999) Urine as anotherpotential source for template DNA in polymerasechain reaction (PCR). American Journal ofPrimatology 48, 299–304

Hearn JP, Lunn SF, Burden FJ, Pilcher MM (1975)Management of marmosets for biomedicalresearch. Laboratory Animals 9, 125–34

Hester HR, Essex HE, Mann FC (1940) Secretion ofurine in the chicken (Gallus domesticus).American Journal of Physiology 128, 592

Home Office (1989) Code of practice for the housingand care of animals used in scientific procedures.In: Animals (Scientific procedures) Act 1986. HerMajesty’s Stationery Office, London, UK

Hubrecht R (2002) Comfortable quarters for cats inresearch institutions. In: Comfortable Quartersfor Laboratory Animals, 9th edn (Reinhardt V,Reinhardt A, eds). Washington DC: AnimalWelfare Institute

Hughes BO, Black AJ (1976) The influence of han-dling on egg production, egg shell quality andavoidance behaviour of hens. British PoultryScience 17, 135–44

Jackson AJ, Sutherland JC (1984) Novel device forquantitatively collecting small volumes of urinefrom laboratory rats. Journal of PharmaceuticalSciences 73, 816–18

Jennings M, Batchelor GR, Brain PF, Dick A, Elliot H,Francis RJ, Hubrecht RC, Hurst JL, Morton DB,Peters AG, Raymond R, Sales GD, Sherwin CM,West C (1998) Refining rodent husbandry: themouse. Report of the Rodent Refinement WorkingParty. Laboratory Animals 32, 233–59

Jezierski TA, Konecka AM (1996) Handling andrearing results in young rabbits. Applied AnimalBehaviour Science 46, 243–50

Jones RB, Dilks RA, Nowell NW (1973) A methodfor the collection of individual mouse urine.Physiology and Behavior 10, 163–4

Kartchner RJ, Rittenhouse LR (1979) A faeces-urineseparator for making total fecal collections fromthe female bovine. Journal of Range Management32, 404–5

Kelley TM, Bramblett CA (1981) Urine collectionfrom vervet monkeys by instrumental condition-ing. Americal Journal of Primatology 1, 95–7

Khosho FK, Kaufmann RC, Amankwah KS (1985) Asimple and efficient method for obtaining urinesamples from rats. Laboratory Animal Science35, 513–14

Klausner JS, Osborne CA, Hilgren J, et al. (1975) Theinterpretation and misinterpretation of bacteriuria.Minnesota Veterinarian 15, 43–7

Kohn CW, Strasser SL (1986) 24-hour renal clearanceand excretion of endogenous substances in themare. American Journal of Veterinary Research47, 1332–7

Kruger JM, Osborne CA, Ulrich LK (1996)Cystocentesis. Diagnostic and therapeutic consid-erations. The Veterinary Clinics of NorthAmerica. Small Animal Practice 26, 353–61

Kurien BT, Scofield RH (1999) Mouse urine collec-tion using clear plastic wrap. Laboratory Animals33, 83–6

Lane VM, Merritt AM (1983) Reliability of single-sample phosphorus fractional excretion determi-nation as a measure of daily phosphorus renalclearance in equids. American Journal ofVeterinary Research 44, 500–2

Lartigue CW, Driscoll TB, Johnson PC (1978)Low-temperature urine collection apparatus forlaboratory rodents. Laboratory Animal Science28, 594–7

Lawlor MM (2002) Comfortable quarters for rats inresearch institutions. In: Comfortable Quartersfor Laboratory Animals, 9th edn (Reinhardt V,Reinhardt A, eds). Washington DC: AnimalWelfare Institute

Lensink BJ, Boivin X, Pradel P, Le Neindre P, Veissier I(2000) Reducing veal calves’ reactivity to peopleby providing additional human contact. Journal ofAnimal Science 78, 1213–18

Levine S (1985) A definition of stress? In: AnimalStress (Moberg GP, ed). Baltimore, MD: WaverlyPress, pp 51–69

Lopez-Anaya A, Unadkat JD, Schumann LA (1990)Simple and effective procedure for complete urinecollection from infant macaques (Macacanemestrina). Journal of Pharmacological Methods24, 105–9

Lunn, SF (1989) Systems for collection of urine inthe captive common marmoset, Callithrixjacchus. Laboratory Animals 23, 353–6

Manser CE, Broom DM, Overend P, Morris TH(1998) Investigation into the preference of labora-tory rats for nest-boxes and nesting materials.Laboratory Animals 32, 23–35

Manser CE, Elliott H, Morris TH, Broom DM (1996)The use of a novel operant test to determine thestrength of preference for flooring in laboratoryrats. Laboratory Animals 30, 1–6

Manser CE, Morris TH, Broom DM (1995) An inves-tigation into the effects of solid or grid cageflooring on the welfare of laboratory rats.Laboratory Animals 29, 353–63

Matandos CK, Franz DR (1980) Collection of urinefrom caged laboratory cats. Laboratory AnimalScience 30, 562–4

Mayrs EB (1924) Secretion as a factor in eliminationby the bird’s kidney. Journal of Physiology 58, 276

Merkenschlager M, Wilk W, eds (1979)Recommendation for the keeping of laboratoryanimals in accordance with animal protectionprinciples. In: Recommendations for the PossibleLimitation and Substitution of Experiments withAnimals. Berlin: Paul Parey

Miller LC, Bard KA, Juno CJ, Nadler RD (1986)Behavioral responsiveness of young chimpanzees(Pan troglodytes) to a novel environment. FoliaPrimatologica (Basel) 47, 128–42

Milroy T (1903) Journal of Physiology 30, 47Minkowski O (1886) Arch. F. Exper. Path. u.

Pharmakol 21, 41Morrisey JK, Ramer JC (1999) Ferrets. Clinical

pathology and sample collection. The VeterinaryClinics of North America. Exotic Animal Practice2, 553–64, vi: Review

Morton DB (2002) Foreword. In: ComfortableQuarters for Laboratory Animals, 9th edn(Reinhardt V, Reinhardt A, eds). Washington DC:Animal Welfare Institute

Mulcahy JJ, Baehler RW, Malvin RL (1978) Acystostomy cannula for dogs. InvestigativeUrology 16, 33–4

Murray MJ (1997) Diagnostic techniques in avianmedicine. Seminars in Avian Exotic Pet Medicine6, 48

Musacchia XJ, Deavers DR, Meininger GA, Davis TP(1980) A model for hypokinesia: effects on muscleatrophy in the rat. Journal of Applied Physiology48, 479–86

National Research Council (1996) Guide for theCare and Use of Laboratory Animals. WashingtonDC: National Academy Press

National Research Council (1998) The PsychologicalWell-Being of Nonhuman Primates. WashingtonDC: National Academy Press

Ness RD (1999) Clinical pathology and samplecollection of exotic small mammals. TheVeterinary Clinics of North America. ExoticAnimal Practice 2, 591–620, vi: Review

Pastoor FJ, van’t Klooster AT, Beynen AC (1990) Analternative method for the quantitative collectionof faeces and urine of cats as validated by thedetermination of mineral balance. Zeitschrift furVersuchstierkunde 33, 259–63

Paton DN (1910) Journal of Physiology 39, 485Paulson GD, Cottrell DJ (1984) An apparatus for

quantitative collection of urine from male cattle,sheep, and swine. American Journal of VeterinaryResearch 45, 2150–1

Peacock AC, Harris RS (1950) Plastic house forthe quantitative separation of urine and faecesexcreted by male rats. Archives of Biochemistryand Biophysics 27, 198–201

Perline IR (1971) An inexpensive mouse urinecollection system. Physiology and Behaviour6, 597

Poole TB (1997) Happy animals make good science.Laboratory Animals 31, 116–24

Rawlings CA, Bisgard GE (1975) Renal clearance andexcretion of endogenous substances in the smallpony. American Journal of Veterinary Research36, 45–8

Reinhardt V, Reinhardt A (2000a) Blood collectionprocedure of laboratory primates: a neglected



variable in biomedical research. Journal of AppliedAnimal Welfare Science 4, 143–9

Reinhardt V, Reinhardt A (2000b) Social enhancementfor adult nonhuman primates in research laborato-ries: a review. Name of journal please 29, 34–41

Reinhardt V (2002) Comfortable quarters for nonhu-man primates in research institutions. In:Comfortable Quarters for Laboratory Animals,9th edn (Reinhardt V, Reinhardt A, eds).Washington DC: Animal Welfare Institute

Rochlitz I (2000) Recommendations for the housingand care of domestic cats in laboratories.Laboratory Animals 34, 1–9

Rochlitz I (2002) Comfortable quarters for cats inresearch institutions. In: Comfortable Quartersfor Laboratory Animals, 9th edn (Reinhardt V,Reinhardt A, eds). Washington DC: AnimalWelfare Institute

Russel FG, Wouterse AC, Hekman P, Grutters GJ,van Ginneken CA (1987) Quantitative urinecollection in renal clearance studies in the dog.Journal of Pharmacological Methods 17, 125–36

Russell WMS (2002) The ill effects of uncomfortablequarters. In: Comfortable Quarters for LaboratoryAnimals, 9th edn (Reinhardt V, Reinhardt A, eds).Washington DC: Animal Welfare Institute

Sharpe NC (1912) On the secretion of urine in birds.American Journal of Physiology 31, 75

Sherwin CM (2002) Comfortable quarters for mice inresearch institutions. In: Comfortable Quartersfor Laboratory Animals, 9th edn (Reinhardt V,Reinhardt A, eds). Washington DC: AnimalWelfare Institute

Smith CR, Felton JS, Taylor RT (1981) Description ofa disposable individual-mouse urine collectionapparatus. Laboratory Animal Science 31, 80–2

Snowdon CT, Savage A, McConnell PB (1985) Abreeding colony of cotton-top tamarins (Saguinusoedipus). Laboratory Animal Science 35, 477–80

Stone AM, et al. (1996) Positive reinforcementtraining for voluntary movement of group-housedchimpanzees (Abstracts). XVIth Congress of theInternational Primatological Society/XIXthConference of the American Society ofPrimatologists. Madison, Wisconsin, No. 679

Styles DK, Phalen DN (1998) Clinical avian urology.Seminars in Avian Exotic Pet medicine 7, 104

Sunderman FW (1944) Method of collecting albinorat urine. American Journal of Clinical Pathology9, 11–12

Tasker JB (1966) Fluid and electrolyte studies in thehorse. II. An apparatus for the collection of totaldaily urine and faeces from horses. CornellVeterinarian 56, 77–84

Townsend P (1997) Use of in-cage shelters by labora-tory rats. Animal Welfare 6, 95–103

Toon S, Rowland M (1981) A simple restrainingdevice for chronic pharmacokinetic and metabolicstudies in rats. Journal of PharmacologicalMethods 5, 321–3



Van den Berg JS (1996) Modified apparatus for collec-tion of free-flow urine from mares. Journal of theSouth African Veterinary Association 67, 214–16

Van der Noot GW, Fonnesbeck PV, Lydman RK(1965) Equine metabolic stall and collectionharness. Journal of Animal Science 24, 691–6

Van Metre DC, Angelos SM (1999) Miniature pigs.The Veterinary Clinics of North America. ExoticAnimal Practice 2, 519–37, v: Review

Watts RH (1971) A simple capillary tube method forthe determination of the specific gravity of 25 and50 micro l quantities of urine. Journal of ClinicalPathology 24, 667–8

West RW, Stanley JW, Newport GD (1978) Single-mouse urine collection and pH monitoringsystem. Laboratory Animal Science 28, 343–5

White WA (1971) A technique for urine collectionfrom anesthetized male rats. Laboratory AnimalScience 21, 401–2

Wronski TJ, Morey-Holton ER (1987) Skeletalresponse to simulated weightlessness: a compari-son of suspension techniques. Aviation, Space,and Environmental Medicine 58, 63–8

Wurbel H (2001) Ideal homes? Housing effects onrodent brain and behaviour. Trends in Neuro-sciences 24, 207–11

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