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A container for transporting small laboratory animals for magnetic resonance imaging
Journal of Neuroscience Methods 144 (2005) 143–146
Short communication
A container for transporting small laboratory animals for magnetic resonance imaging Hua Zhana , Tsuyoshi Tadaa,∗ , Eiji Fujikurab , Kiyoshi Matsumotoc , Yuichiro Tanakaa , Kazuhiro Hongoa a
Department of Neurosurgery, Shinshu University School of Medicine, 3-1-1 Asahi, Matsumoto 390-8621, Japan b MRI Room, Aizawa Hospital, Matsumoto, Japan c Institute of Experimental Animals, Shinshu University School of Medicine, Matsumoto, Japan Received 15 October 2003; received in revised form 14 October 2004; accepted 14 October 2004
Abstract We have constructed a simple container, consisting of a propylene tube, a High-Efficiency Particulate Aerosol (HEPA) filter and a rubber glove, for transporting small animals to magnetic resonance imaging (MRI) facilities that are located outside a pathogen-free environment. Results of pathogens analysis indicate that the container is able to prevent infection by several microorganisms. The quality of the MR images of mice and rats transported in, and imaged while in the container was satisfactory. This container can be useful for examinations that required the use of instruments located outside clean animal units, ensuring safety for both humans and animals. © 2004 Elsevier B.V. All rights reserved. Keywords: Container; Laboratory animal; MRI; Mouse; Rat; Isolation
1. Introduction Transgenic animal models have become a powerful research tool, especially in molecular biology and medicine (Finn et al., 2003; Correia-Pinto et al., 2002). These and other laboratory animals are bred in special facilities designed which often provide highly clean, pathogen-free conditions (Kanzaki et al., 2001). The lack of sophisticated and expensive instrumentation within these animal units often requires the transport to an animal to a different facility to carry out the required experimental procedure, and the consequence impossibility to return it to the pathogenfree unit (Richmond and Nesby-O’Dell, 2003; Kowalski et al., 2002). One example of this situation is the need to perform magnetic resonance (MR) or computed tomographic (CT) imaging of experimental animals, since there are still very few pathogen-free animal units that can afford this expensive in∗
Corresponding author. Tel.: +81 263 37 2690; fax: +81 263 37 0480. E-mail address: [email protected] (T. Tada).
0165-0270/$ – see front matter © 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.jneumeth.2004.10.008
strumentation. Often, MR or CT systems that are located in a hospital environment are, therefore used. This, in turn raises two major problems: (i) the MR or CT environment needs to remain as clean (and potentially pathogen-free) as the animal unit, and (ii) the large difference in the size between the human and animal area of interest. To solve these problems, we have now designed a simple container for transporting mice and rats between an animal facility and a hospital environment. We report that this container appears to maintain the pathogen-free condition of the transported animal and to allow good quality of MRI images in anaesthetized animals.
2. Materials and methods 2.1. Container A 2-l clear polystyrene bottle was used as the body of the container (Fig. 1). A mouse folder made from half of a 50-ml polypropylene conical tube (Becton Dickinson and
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H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
Fig. 1. Schematic drawing of the procedure for constructing the container. A1 and A2: Cut off the tapered end of the bottle and cut a window (size, about 4 cm × 15 cm) in its side. B1 and B2: Cut a 50-ml tube in half. C: Lateral (left) and coronal (right) views of the finished container, after fixing the tube (B2) in a suitable position within the bottle (A2). D: Lateral (left) and coronal (right) photographs of the container. E: Placing a hand in the rubber glove, the animal can be positioned as required without opening the container.
Company, Franklin Lakes, NJ) was fixed to the bottom of the container. Because most pathogens are contracted via the airborne route, the opening of the bottle (i.e., the respiratory window) was covered with a High-Efficiency Particulate Aerosol (HEPA) filter (Havenaar et al., 1993). The containers and the carrying box were sterilized with ethylene oxide gas before use.
2.2. Animals All experiments were carried out in accordance with national and local ethical committee guidelines. Four-week-old male C57BL/6 mice and 13-week-old male Wister rats were purchased from SLC Inc. (Shizuoka, Japan), and were housed in the Shinshu University Animal House.
H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
145
2.3. Sedation of animals Animals used in this study were housed in a non-pathogen free area. They were sedated by intraperitoneal injection of a mixture of ketamine hydrochloride (50 mg/kg body weight) and medetomidine (10 mg/kg body weight). To reverse the anesthesia, antipamezole hydrochloride was injected at 1–2.5 mg/kg (Cruz et al., 1998). The head of each animal was fixed in the sterilized container in supine position, and the open part of the container was closed using a sterilized surgical rubber glove. A wrong positioning of the animal, with respect to the intended plane of imaging, could be easily corrected using the rubber glove (Fig. 1E). 2.4. Magnetic resonance imaging The MRI imaging was performed with a 1.5-T wholebody MR imager (Signa Advantage, General Electric, Milwaukee, Wis.), using standard gradient hardware with a round surface coil (diameter: 3 inches). T2-weighted images were obtained in sequences: coronal plane; slice, 0.5 cm; TR/TE = 2000/90 ms; FOV = 12 mm × 6 mm; data matrix = 256 × 192 points. For T1-weighted MRI, TR/TE=600/11.9 ms, FOV = 12 mm × 6 mm, and data matrix = 256 × 256 points. To improve the imaging, 2 bottles, each containing 500 ml of isotonic sodium chloride solution, were placed on both sides of the container.
Fig. 2. A: T2-weighted coronal MR image of a mouse head. It is easy to distinguish the cortex, lateral ventricle, dorsal third ventricle, brain stem, and third ventricle. Also identifiable are the caudate putamen, lateral globus pallidus, basal nucleus, and reticular thalamic nucleus. B: T1-weighted coronal MR image of a rat head.
2.5. Pathogen analysis
4. Discussion
Five mice transported in the container were microbiologically monitored by culture, microscopy and serology analyses (Mercian Cleantec Corporation, Kanagawa, Japan). They were free of the following organisms: Pseudomonas aeruginosa, Pasteurella pneumotropica, Bordetella bronchiseptica, Salmonella spp., Corynebacterium kutscheri, Tyzzer’s organism (Clostridium piliformis), Dermatophytes, Mycoplasma pulmonis, Hemagglutinating virus of Japan, Mouse hepatitis virus, Giardia spp., Spironucleus muris and Syphacia spp (Itoh, 1999).
It is difficult to transport laboratory animals out of clean, pathogen-free units, into areas for experimentation that are located in a non-pathogen-free facility. In addition, once the animal has been removed from a clean unit cannot be returned, as they have become potential carriers of diseases (Lane-Petter, 1976). Indeed, the ability to remove animals from clean animal units and to keep them isolated under clean conditions during experimentation, would then allow researchers to dramatically decrease the number of animals used. There are many kinds of containers for transporting laboratory animals (Townsend and Robinson, 1976). Most are made from wood or paper, and use for in transporting animals from breeders to animal research facilities. We have constructed a container for use with MRI, which requires it to be of a non-magnetic material that keeps the animal in a dust and pathogen-free environment. Compact High-Efficiency Particulate Aerosol (HEPA) filter systems are commonly used to reduce airborne dust concentrations (Robert and Demers, 2001; Abraham, 1999; Willeke et al., 2001). A HEPA-grade device is defined as any filter capable of trapping at least 99.97% of particles with a diameter of 0.3 m. Phillpotts et al. (1997) kept
3. Results No problem was encountered during any of the procedures involved in the insertion or repositioning of the animal in the container. The results of the pathogen analysis were negative indicating that the container was able to prevent infection of all the assayed microorganisms (see methods). As shown in Fig. 2, the quality of a rat and a mouse MR image was of satisfactory quality. The cortex, lateral ventricle, dorsal third ventricle and brain stem were easily distinguishable (Paxinos and Franklin, 2001).
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H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
animals in a chamber closed by a HEPA filter, and exposed them to a pathogen aerosolized using a Collison nebulizer. Although our animals were housed in a non-pathogen-free zone for more than 4 weeks, the pathogen analysis showed that the HEPA filter prevented infection by the assayed microorganisms. MRI technology has enabled clear imaging of small laboratory animals, including embryos (Schneider et al., 2003; Redwine et al., 2003; Dhenain et al., 2001; Choi et al., 2003). Because of their small body weight, however, it is difficult to obtain clear images of small laboratory animals using a hospital MRI system. Our container allowed good quality imaging of mice and rats. This was also improved by placing a bottle of isotonic sodium chloride solution on each side of the container to prevent disturbance in MRI analysis. Furthermore, by combining a non-magnetic mouse head holder for stereotaxic surgery (Tada et al., 2002) with the present container, pre- and post-operative images can be easily obtained.
References Abraham ME. Microanalysis of indoor aerosols and the impact of a compact high-efficiency particulate air (HEPA) filter system. Indoor Air 1999;9(1):33–40. Choi IY, Lee SP, Guilfoyle DN, Helpern JA. In vivo NMR studies of neurodegenerative diseases in transgenic and rodent models. Neurochem Res 2003;28(7):987–1001. Correia-Pinto J, Henriques-Coelho T, Oliveira SM, Moreira AF. Evaluation of biventricular function in the rat: a new experimental model. Rev Port Cardiol 2002;21(11):1295–302. Cruz JI, Loste JM, Burzaco OH. Observations on the use of medetomidine/ketamine and its reversal with atipamezole for chemical restraint in the mouse. Lab Anim 1998;32(1):18–22. Dhenain M, Ruffins SW, Jacobs RE. Three-dimensional digital mouse atlas using high-resolution MRI. Dev Biol 2001;232:458–70. Finn DA, Rutledge-Gorman MT, Crabbe JC. Genetic animal models of anxiety. Neurogenetics 2003;4(3):109–35. Havenaar R, Meijer JC, Morton DB, Ritskes-Hoitinga J, Zwart P. Biology and husbandry of laboratory animals. In: Ven Zutphen LFM,
Baumans V, Beynen AC, editors. Principles of Laboratory Animal Science. Amsterdam: Elsevier; 1993. p. 17–74. Itoh T. Quality testing system for SPF animals in Japan and problems in the management of such systems. Institute for Laboratory Animal Research. Microbial and phenotypic definition of rats and mice: proceeding of the 1998 US/Japan conference. The National Academies Press, 1999: 15–23. Kanzaki M, Fujieda M, Furukawa T. Effects of suspension of airconditioning on airtight-type racks. Exp Anim 2001;50(5):379–85. Kowalski WJ, Bahnfleth WP, Carey DD. Engineering control of airborne disease transmission in animal laboratories. Contemp Top Lab Anim Sci 2002;41(3):9–17. Lane-Petter W. The laboratory mouse. UFAW. The UFAW handbook on the care and management of laboratory animals. Longman Group Limited: New York, 1976: 193–209. Paxinos G, Franklin KBJ. The mouse brain in stereotaxic coordinates. 2nd ed. Son Diego: Academic Press; 2001. Phillpotts RJ, Brooks TJ, Cox CS. A simple device for the exposure of animals to infectious microorganisms by the airborne route. Epidemiol Infect 1997;118(1):71–5. Redwine JM, Kosofsky B, Jacobs RE, Games D, Reilly JF, Morrison JH, Young WG, Bloom FE. Dentate gyrus volume is reduced before onset of plaque formation in PDAPP mice: a magnetic resonance microscopy and stereologic analysis. PNAS 2003;100(3):1381–6. Richmond JY, Nesby-O’Dell S. Biosecurity for animal facilities and associated laboratories. Lab Anim 2003;32(1):32–5. Robert R, Demers BS. RRT. Bacterial/viral filtration, let the breather beware. Chest 2001;120:1377–89. Schneider JE, Bamforth SD, Farthing CR, Clarke K, Neubauer S, Bhattacharya S. High-resolution imaging of normal anatomy, and neural and adrenal malformations in mouse embryos using magnetic resonance microscopy. J Anat 2003;202:239–47. Tada T, Wendland M, Watson N, Kuriyama N, Kuriyama H, Roberts T, Burns M, Weiss W, Israel MA. A head holder for magnetic resonance imaging that allows the stereotaxic alignment of spontaneously occurring intracranial mouse tumors. J Neurosci Methods 2002;116: 1–7. Townsend GH, Robinson MH. Transport of laboratory animals In UFAW, editor. The UFAW handbook on the care and management of laboratory animals. Longman Group Limited: New York, 1976: 106– 116. Willeke K, Trakumas S, Grinshpun SA, Reponen T, Trunov M, Friedman W. Test methods for evaluating the filtration and particulate emission characteristics of vacuum cleaners. AIHAJ 2001;62(3):313–21.
Short communication
A container for transporting small laboratory animals for magnetic resonance imaging Hua Zhana , Tsuyoshi Tadaa,∗ , Eiji Fujikurab , Kiyoshi Matsumotoc , Yuichiro Tanakaa , Kazuhiro Hongoa a
Department of Neurosurgery, Shinshu University School of Medicine, 3-1-1 Asahi, Matsumoto 390-8621, Japan b MRI Room, Aizawa Hospital, Matsumoto, Japan c Institute of Experimental Animals, Shinshu University School of Medicine, Matsumoto, Japan Received 15 October 2003; received in revised form 14 October 2004; accepted 14 October 2004
Abstract We have constructed a simple container, consisting of a propylene tube, a High-Efficiency Particulate Aerosol (HEPA) filter and a rubber glove, for transporting small animals to magnetic resonance imaging (MRI) facilities that are located outside a pathogen-free environment. Results of pathogens analysis indicate that the container is able to prevent infection by several microorganisms. The quality of the MR images of mice and rats transported in, and imaged while in the container was satisfactory. This container can be useful for examinations that required the use of instruments located outside clean animal units, ensuring safety for both humans and animals. © 2004 Elsevier B.V. All rights reserved. Keywords: Container; Laboratory animal; MRI; Mouse; Rat; Isolation
1. Introduction Transgenic animal models have become a powerful research tool, especially in molecular biology and medicine (Finn et al., 2003; Correia-Pinto et al., 2002). These and other laboratory animals are bred in special facilities designed which often provide highly clean, pathogen-free conditions (Kanzaki et al., 2001). The lack of sophisticated and expensive instrumentation within these animal units often requires the transport to an animal to a different facility to carry out the required experimental procedure, and the consequence impossibility to return it to the pathogenfree unit (Richmond and Nesby-O’Dell, 2003; Kowalski et al., 2002). One example of this situation is the need to perform magnetic resonance (MR) or computed tomographic (CT) imaging of experimental animals, since there are still very few pathogen-free animal units that can afford this expensive in∗
Corresponding author. Tel.: +81 263 37 2690; fax: +81 263 37 0480. E-mail address: [email protected] (T. Tada).
0165-0270/$ – see front matter © 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.jneumeth.2004.10.008
strumentation. Often, MR or CT systems that are located in a hospital environment are, therefore used. This, in turn raises two major problems: (i) the MR or CT environment needs to remain as clean (and potentially pathogen-free) as the animal unit, and (ii) the large difference in the size between the human and animal area of interest. To solve these problems, we have now designed a simple container for transporting mice and rats between an animal facility and a hospital environment. We report that this container appears to maintain the pathogen-free condition of the transported animal and to allow good quality of MRI images in anaesthetized animals.
2. Materials and methods 2.1. Container A 2-l clear polystyrene bottle was used as the body of the container (Fig. 1). A mouse folder made from half of a 50-ml polypropylene conical tube (Becton Dickinson and
144
H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
Fig. 1. Schematic drawing of the procedure for constructing the container. A1 and A2: Cut off the tapered end of the bottle and cut a window (size, about 4 cm × 15 cm) in its side. B1 and B2: Cut a 50-ml tube in half. C: Lateral (left) and coronal (right) views of the finished container, after fixing the tube (B2) in a suitable position within the bottle (A2). D: Lateral (left) and coronal (right) photographs of the container. E: Placing a hand in the rubber glove, the animal can be positioned as required without opening the container.
Company, Franklin Lakes, NJ) was fixed to the bottom of the container. Because most pathogens are contracted via the airborne route, the opening of the bottle (i.e., the respiratory window) was covered with a High-Efficiency Particulate Aerosol (HEPA) filter (Havenaar et al., 1993). The containers and the carrying box were sterilized with ethylene oxide gas before use.
2.2. Animals All experiments were carried out in accordance with national and local ethical committee guidelines. Four-week-old male C57BL/6 mice and 13-week-old male Wister rats were purchased from SLC Inc. (Shizuoka, Japan), and were housed in the Shinshu University Animal House.
H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
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2.3. Sedation of animals Animals used in this study were housed in a non-pathogen free area. They were sedated by intraperitoneal injection of a mixture of ketamine hydrochloride (50 mg/kg body weight) and medetomidine (10 mg/kg body weight). To reverse the anesthesia, antipamezole hydrochloride was injected at 1–2.5 mg/kg (Cruz et al., 1998). The head of each animal was fixed in the sterilized container in supine position, and the open part of the container was closed using a sterilized surgical rubber glove. A wrong positioning of the animal, with respect to the intended plane of imaging, could be easily corrected using the rubber glove (Fig. 1E). 2.4. Magnetic resonance imaging The MRI imaging was performed with a 1.5-T wholebody MR imager (Signa Advantage, General Electric, Milwaukee, Wis.), using standard gradient hardware with a round surface coil (diameter: 3 inches). T2-weighted images were obtained in sequences: coronal plane; slice, 0.5 cm; TR/TE = 2000/90 ms; FOV = 12 mm × 6 mm; data matrix = 256 × 192 points. For T1-weighted MRI, TR/TE=600/11.9 ms, FOV = 12 mm × 6 mm, and data matrix = 256 × 256 points. To improve the imaging, 2 bottles, each containing 500 ml of isotonic sodium chloride solution, were placed on both sides of the container.
Fig. 2. A: T2-weighted coronal MR image of a mouse head. It is easy to distinguish the cortex, lateral ventricle, dorsal third ventricle, brain stem, and third ventricle. Also identifiable are the caudate putamen, lateral globus pallidus, basal nucleus, and reticular thalamic nucleus. B: T1-weighted coronal MR image of a rat head.
2.5. Pathogen analysis
4. Discussion
Five mice transported in the container were microbiologically monitored by culture, microscopy and serology analyses (Mercian Cleantec Corporation, Kanagawa, Japan). They were free of the following organisms: Pseudomonas aeruginosa, Pasteurella pneumotropica, Bordetella bronchiseptica, Salmonella spp., Corynebacterium kutscheri, Tyzzer’s organism (Clostridium piliformis), Dermatophytes, Mycoplasma pulmonis, Hemagglutinating virus of Japan, Mouse hepatitis virus, Giardia spp., Spironucleus muris and Syphacia spp (Itoh, 1999).
It is difficult to transport laboratory animals out of clean, pathogen-free units, into areas for experimentation that are located in a non-pathogen-free facility. In addition, once the animal has been removed from a clean unit cannot be returned, as they have become potential carriers of diseases (Lane-Petter, 1976). Indeed, the ability to remove animals from clean animal units and to keep them isolated under clean conditions during experimentation, would then allow researchers to dramatically decrease the number of animals used. There are many kinds of containers for transporting laboratory animals (Townsend and Robinson, 1976). Most are made from wood or paper, and use for in transporting animals from breeders to animal research facilities. We have constructed a container for use with MRI, which requires it to be of a non-magnetic material that keeps the animal in a dust and pathogen-free environment. Compact High-Efficiency Particulate Aerosol (HEPA) filter systems are commonly used to reduce airborne dust concentrations (Robert and Demers, 2001; Abraham, 1999; Willeke et al., 2001). A HEPA-grade device is defined as any filter capable of trapping at least 99.97% of particles with a diameter of 0.3 m. Phillpotts et al. (1997) kept
3. Results No problem was encountered during any of the procedures involved in the insertion or repositioning of the animal in the container. The results of the pathogen analysis were negative indicating that the container was able to prevent infection of all the assayed microorganisms (see methods). As shown in Fig. 2, the quality of a rat and a mouse MR image was of satisfactory quality. The cortex, lateral ventricle, dorsal third ventricle and brain stem were easily distinguishable (Paxinos and Franklin, 2001).
146
H. Zhan et al. / Journal of Neuroscience Methods 144 (2005) 143–146
animals in a chamber closed by a HEPA filter, and exposed them to a pathogen aerosolized using a Collison nebulizer. Although our animals were housed in a non-pathogen-free zone for more than 4 weeks, the pathogen analysis showed that the HEPA filter prevented infection by the assayed microorganisms. MRI technology has enabled clear imaging of small laboratory animals, including embryos (Schneider et al., 2003; Redwine et al., 2003; Dhenain et al., 2001; Choi et al., 2003). Because of their small body weight, however, it is difficult to obtain clear images of small laboratory animals using a hospital MRI system. Our container allowed good quality imaging of mice and rats. This was also improved by placing a bottle of isotonic sodium chloride solution on each side of the container to prevent disturbance in MRI analysis. Furthermore, by combining a non-magnetic mouse head holder for stereotaxic surgery (Tada et al., 2002) with the present container, pre- and post-operative images can be easily obtained.
References Abraham ME. Microanalysis of indoor aerosols and the impact of a compact high-efficiency particulate air (HEPA) filter system. Indoor Air 1999;9(1):33–40. Choi IY, Lee SP, Guilfoyle DN, Helpern JA. In vivo NMR studies of neurodegenerative diseases in transgenic and rodent models. Neurochem Res 2003;28(7):987–1001. Correia-Pinto J, Henriques-Coelho T, Oliveira SM, Moreira AF. Evaluation of biventricular function in the rat: a new experimental model. Rev Port Cardiol 2002;21(11):1295–302. Cruz JI, Loste JM, Burzaco OH. Observations on the use of medetomidine/ketamine and its reversal with atipamezole for chemical restraint in the mouse. Lab Anim 1998;32(1):18–22. Dhenain M, Ruffins SW, Jacobs RE. Three-dimensional digital mouse atlas using high-resolution MRI. Dev Biol 2001;232:458–70. Finn DA, Rutledge-Gorman MT, Crabbe JC. Genetic animal models of anxiety. Neurogenetics 2003;4(3):109–35. Havenaar R, Meijer JC, Morton DB, Ritskes-Hoitinga J, Zwart P. Biology and husbandry of laboratory animals. In: Ven Zutphen LFM,
Baumans V, Beynen AC, editors. Principles of Laboratory Animal Science. Amsterdam: Elsevier; 1993. p. 17–74. Itoh T. Quality testing system for SPF animals in Japan and problems in the management of such systems. Institute for Laboratory Animal Research. Microbial and phenotypic definition of rats and mice: proceeding of the 1998 US/Japan conference. The National Academies Press, 1999: 15–23. Kanzaki M, Fujieda M, Furukawa T. Effects of suspension of airconditioning on airtight-type racks. Exp Anim 2001;50(5):379–85. Kowalski WJ, Bahnfleth WP, Carey DD. Engineering control of airborne disease transmission in animal laboratories. Contemp Top Lab Anim Sci 2002;41(3):9–17. Lane-Petter W. The laboratory mouse. UFAW. The UFAW handbook on the care and management of laboratory animals. Longman Group Limited: New York, 1976: 193–209. Paxinos G, Franklin KBJ. The mouse brain in stereotaxic coordinates. 2nd ed. Son Diego: Academic Press; 2001. Phillpotts RJ, Brooks TJ, Cox CS. A simple device for the exposure of animals to infectious microorganisms by the airborne route. Epidemiol Infect 1997;118(1):71–5. Redwine JM, Kosofsky B, Jacobs RE, Games D, Reilly JF, Morrison JH, Young WG, Bloom FE. Dentate gyrus volume is reduced before onset of plaque formation in PDAPP mice: a magnetic resonance microscopy and stereologic analysis. PNAS 2003;100(3):1381–6. Richmond JY, Nesby-O’Dell S. Biosecurity for animal facilities and associated laboratories. Lab Anim 2003;32(1):32–5. Robert R, Demers BS. RRT. Bacterial/viral filtration, let the breather beware. Chest 2001;120:1377–89. Schneider JE, Bamforth SD, Farthing CR, Clarke K, Neubauer S, Bhattacharya S. High-resolution imaging of normal anatomy, and neural and adrenal malformations in mouse embryos using magnetic resonance microscopy. J Anat 2003;202:239–47. Tada T, Wendland M, Watson N, Kuriyama N, Kuriyama H, Roberts T, Burns M, Weiss W, Israel MA. A head holder for magnetic resonance imaging that allows the stereotaxic alignment of spontaneously occurring intracranial mouse tumors. J Neurosci Methods 2002;116: 1–7. Townsend GH, Robinson MH. Transport of laboratory animals In UFAW, editor. The UFAW handbook on the care and management of laboratory animals. Longman Group Limited: New York, 1976: 106– 116. Willeke K, Trakumas S, Grinshpun SA, Reponen T, Trunov M, Friedman W. Test methods for evaluating the filtration and particulate emission characteristics of vacuum cleaners. AIHAJ 2001;62(3):313–21.