Generation of Mammalian Host-adapted Leptospira interrogans by Cultivation in Peritoneal Dialysis Membrane Chamber Implantation in Rats

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PLOS Pathogens
Mar 2014



Leptospira interrogans can infect a myriad of mammalian hosts, including humans (Bharti et al., 2003; Ko et al., 2009). Following acquisition by a suitable host, leptospires disseminate via the bloodstream to multiple tissues, including the kidneys, where they adhere to and colonize the proximal convoluted renal tubules (Athanazio et al., 2008). Infected hosts shed large number of spirochetes in their urine and the leptospires can survive in different environmental conditions before transmission to another host. Differential gene expression by Leptospira spp. permits adaption to these new conditions. Here we describe a protocol for the cultivation of Leptospira interrogans within Dialysis Membrane Chambers (DMCs) implanted into the peritoneal cavities of Sprague-Dawley rats (Caimano et al., 2014). This technique was originally developed to study mammalian adaption by the Lyme disease spirochete, Borrelia burgdorferi (Akins et al., 1998; Caimano, 2005). The small pore size (8,000 MWCO) of the dialysis membrane tubing used for this procedure permits access to host nutrients but excludes host antibodies and immune effector cells. Given the physiological and environmental similarities between DMCs and the proximal convoluted renal tubule, we reasoned that the DMC model would be suitable for studying in vivo gene expression by L. interrogans. In a 20 to 30 min procedure, DMCs containing virulent leptospires are surgically-implanted into the rat peritoneal cavity. Nine to 11 days post-implantation, DMCs are explanted and organisms recovered. Typically, a single DMC yields ~109 mammalian host-adapted leptospires (Caimano et al., 2014). In addition to providing a facile system for studying the transcriptional and physiologic changes pathogenic L. interrogans undergo within the mammal, the DMC model also provides a rationale basis for selecting new targets for mutagenesis and the identification of novel virulence determinants.
Caution: Leptospira interrogans is a BSL-2 level pathogen and known to be excreted in the urine of infected animals. Animals should be handled and disposed of using recommended Animal Biosafety Levels (ABSL) for infectious agents using vertebrate animal guidelines.
Note: All protocols using live animals must conform to governmental regulations regarding the care and use of laboratory animals. The success of this protocol is dependent on the proper use of aseptic techniques during all stages of both dialysis membrane chamber preparation and animal surgery.

Keywords: Spirochetes (螺旋体), Leptospirosis (钩端螺旋体病), Mammalian host adaptation (哺乳动物宿主适应性), Gene expression (基因的表达), Virulence (毒力)

Materials and Reagents

  1. Adult female Sprague-Dawley rats (175-200 g)
  2. EMJH medium (prepared as described in Supplementary data)
  3. Bovine serum albumin (BSA)
    EMJH+BSA (10 mg/ml BSA, BSA concentration 20 mg/ml) (Millipore, catalog number: 810037 )
    Note: The quality of the albumin used for the cultivation of leptospires is critical. Please verify that the albumin used for DMC implantation supports the growth of virulent leptospires under standard in vitro growth conditions. Other sources of BSA were tested (e.g., Millipore, catalog number: 840644 and Sigma-Aldrich, catalog number: A-9647 ) with similar results.
  4. Leptospira interrogans, logarithmic phase, grown under standard in vitro conditions (28-30 °C) (Zuerner, 2005)
  5. Ultrapure water (deionized, distilled) (e.g., Milli-Q)
  6. 1 mM EDTA (pH 8.0)
  7. Regenerated cellulose dialysis membrane tubing (SpectrumTM Spectra/PorTM 1 RC, 6,000 to 8,000 MWCO, 32 mm width) (Spectrum Labs, catalog number: 132655 )
  8. Filter units (0.22 µm, 250 ml, sterile) (e.g., Millipore Stericup® filter unit)
  9. Ketamine/xylazine anesthetic cocktail (40-80 mg per kg/ 5-10 mg per kg, administered intramuscularly)
  10. Ophthalmic ointment (e.g., Puralube)
  11. Betadine® surgical scrub solution
  12. Carprofen (5-10 mg per kg, administered subcutaneously)
  13. Ethanol (70%)
  14. Ketamine/xylazine anesthetic cocktail (see Recipes)
  15. EMJH + BSA medium (see Recipes)


  1. Surgical gloves (individually wrapped, sterile)
  2. Disposable serological pipets (10 ml sterile, individually-wrapped)
  3. Surgical drape (cut into 12 in (46 cm) squares, sterile)
  4. Gauze (4 x 4 in sterile)
  5. Microscope equipped with a dark field condenser (e.g., Olympus, model: BX40 )
  6. Three 2-L Pyrex beakers (each containing a magnetic stir bar)
  7. Hot plate with stirring option
  8. Extra-long blunt end forceps (sterile)
  9. Pipet filler (e.g., Drummond Scientific, model: Pipet-Aid )
  10. Surgical instrument pack, one per animal, sterilized prior to use and kept within sterile package:
    Scalpel blades (No. 10)
    Scalpel blade holder
    Scissors, iris, 4 in (~10.2-cm)
    Tissue forceps, 5 ½ in (~14 cm) (1x 2-tooth dissecting or Adson-Brown)
    Tissue forceps, 5 ½ in (~14 cm) (blunt-end)
    Needle holder forceps with built-in scissors (e.g., Olsen Hegar) (5 ½ in)
    (~14 cm)
    Suture, Ethicon 40, SH 1, 27 in (~68.6 cm) coated Vicryl, violet-braided suture
    Autoclip® 9-mm stainless steel wound closure clips and applicator
  11. Biological safety cabinet (Biosafety Level 2, model: BSL2 )
  12. Circulating warm-water blanket and pump
  13. Electric hair/fur clippers
  14. Glass bead sterilizer (e.g., Braintree Scientific, model: Germinator 500 ) (optional)


  1. Preparation of sterile dialysis membrane tubing
    1. Fill three 2-Liter Pyrex beakers as follows:
      Beaker 1: 1 - 1.5 L Ultrapure water
      Beaker 2: 1 - 1.5 L 1 mM EDTA (pH 8.0)
      Beaker 3: 1 - 1.5 L Ultrapure water
    2. Add a magnetic stir bar to each beaker, cover with heavy-duty aluminum foil and autoclave.
      Note: To ensure sterility, beakers should be autoclaved on the same day at the procedure.
    3. Place each beaker on hot plate and bring to a rolling boil with constant stirring.
    4. Wearing sterile gloves, cut dialysis membrane tubing into strips 7-9 inches (~18-23 cm) in length using sterile scissors. Cut one strip per animal plus 1-2 extra.
    5. Gently tie off one end of the tubing using a simple overhand knot by forming a loop at the top of the DMC and passing the free end of the tubing through the loop. Trim away excess tubing from the tied end using sterile scissors.
    6. Place the tied tubing into Beaker 1 and replace the aluminum foil cover. Be sure to keep the foil cover loose enough to allow steam to escape. Boil tubing for 20 min. with constant stirring.
      Note: Tubing should remain submerged at all times.
    7. Using sterile extra-long forceps, transfer tubing to Beaker 2. Replace aluminum foil cover and boil tubing for 20 min. with constant stirring.
    8. Repeat step A7 using Beaker 3.
    9. Once cool to the touch, transfer Beaker 3 to a Biological safety cabinet (BSC). To ensure sterility, the beaker containing the tubing should remain in the BSC.
      Note: Dialysis membrane tubing may be prepared several days in advance and stored at 4 °C. After boiling, transfer dialysis membrane tubing to the bottom portion of a 0.22 µm Stericup® Filter Unit (Millipore) using sterile blunt tip forceps. Replace the filter unit top and filter in ~100-200 ml of Ultrapure-Q water from Beaker 3. Seal the bottom portion of the filter unit using the sterile cap provided by the manufacturer. The container should be opened only within the BSC.

  2. Preparation of leptospires
    1. Use a standard low passage in vitro culture of virulent L. interrogans, kept at 28-30 °C in EMJH medium, before it reaches stationary phase in the growth curve.
    2. Just prior to starting the surgical procedure, count bacteria by dark field microscopy.
    3. Dilute culture to 106 organisms/ml using fresh EMJH medium.
    4. Working in the BSC, transfer 10 ml of EMJH+BSA to sterile 50-ml conical tubes. Prepare one conical tube per DMC.
      Note: It is important that the EMJH medium used for DMCs is supplemented with additional BSA to maintain the appropriate osmotic pressure.
    5. Add 0.1 ml of dilute culture to each 50-ml conical tube containing EMJH+BSA medium (Final density, 104 organisms/ml).

  3. Preparation of DMCs
    1. Working in the BSC, place several disposable, individually-wrapped serological pipets within the surgical field. Lay down a sterile drape to use as a workspace. Clean the surface of an automatic pipeter using 70% ethanol and place on the sterile drape. After donning a new pair of sterile surgical gloves, loosen the cap of a 50-ml conical tube.
    2. While still wearing gloves, carefully remove a strip of tubing from its storage container using blunt end forceps. Use your free hand to hold the tubing between index finger and thumb. With other hand, remove the cap from the 50-ml conical tube containing diluted bacteria and transfer up to 9 mls to the tubing using a 10-ml disposable serological pipette. While continuing to hold the open end, flatten the unfilled tubing between the fingers of your free hand to eliminate any air bubbles.
    3. Gently twist the top end of the tubing (1-2 turns) to remove any remaining void volume and close off the top of the DMC Use your thumb and index finger to hold on to the twisted area to prevent liquid from leaking out of the DMC while you tie off the top. Tie off the open end using a simple overhand knot (e.g., form a loop at the top of the DMC, passing the free end of the tubing through the loop, then, without releasing your thumb and index finger, gently tighten the knot). Trim away excess tubing from the tied end using sterile scissors.
      Note: Try to make the DMC as taut as possible; this will help to maneuver the chamber into the peritoneal cavity during surgery. The resulting filled DMC should be ~1.5 - 2 in. Larger DMCs may interfere with normal intestinal or bladder functions.
    4. Place the filled DMC into a sterile 50-ml conical tube containing ~5-7 ml of fresh EMJH+ BSA medium.
    5. Repeat steps C3-4 until all bags are filled then proceed directly to the surgical procedure.
      Note: It is not recommended that filled dialysis membrane chambers be stored for extended periods of time as this may increase the risk of contamination.

  4. Peritoneal implantation procedure.
    1. Anesthetize animal by intramuscular injection with the ketamine/xylazine cocktail.
    2. Apply a small amount of ophthalmic ointment to each eye.
    3. Administer preoperative analgesia (e.g., Carprofen).
    4. Shave the abdomen and prepare surgical site (e.g., successive washes with Betadine® Surgical solution followed by alcohol).
    5. Cover the surgical site with sterile gauze and transfer the animal to the BSC in a supine position.
    6. Prior to beginning the procedure, remove the sterile gauze covering the surgical site and perform a ‘toe pinch’ to ensure that the animal is properly anesthetized.
    7. Using a sterile scalpel blade, make a 5 cm incision through the skin only, starting ~2.5 cm below the ribcage, using the xiphoid process as a guide. Using tissue forceps, pull up the skin on either side of the incision and gently trim the fascia connecting the skin to the abdominal wall using a scalpel. Repeat on the other side.
    8. Use the same scalpel, make a small incision (~4 cm) in the abdominal wall, using the linea alba as guide. The incision should be clean and straight.
    9. Using tissue forceps, raise one side of the abdominal incision and place a DMC inside rat peritoneal cavity (Figure 1). Gently push the DMC towards either side of the peritoneal cavity to prevent it from being nicked/ruptured during suturing. Be sure to position so that it does not become entangled in the intestines or interfere with bladder expansion.

    Figure 1. Implantation of DMC into rat peritoneal cavity

    1. Close the abdominal incision site by suturing. Begin by suturing each end of the incision with double knots. Working upwards, close the remaining incision by placing sutures ~2-3 mm apart.
    2. Close the skin incision site using a contiguous line of AutoClip® wound clips (Figure 2). Note: Alternatively, the skin excision site may be closed using subcuticular stitches followed by liquid skin adhesive (e.g., Nexaband).

    Figure 2. Closing the skin incision

    1. Place the rat on top of a clean surgical drape in a clean cage containing fresh bedding. Place the cage on top of a circulating water heating pad to maintain the appropriate core body temperature. Monitor the animal continuously until alert and responsive.
    2. Analgesia should be administered for at least two days post-operatively.
    3. Animals should be monitored at least once daily for the first week post-operatively and then every other day thereafter. If animals show any sign of distress or discomfort, consult institutional veterinary staff immediately.

  5. Recovery of mammalian host adapted organisms from DMCs
    1. At 9 to 12 days after implantation, euthanize animals by CO2 asphyxiation.
    2. Place animal in a BSC in a supine position.
    3. Using sterile surgical scissors, expose the abdominal wall.
    4. Using sterile tissue forceps, lift one side of the sutured abdominal incision site and re-open using sterile scissors.
    5. Locate and remove the DMC using blunt-end forceps. Transfer to a sterile 50-ml conical tube.
    6. Using sterile forceps to hold the DMC by one knot, make a small cut in the tubing just below the knot. Slowly remove the contents of the DMC using a sterile disposable serological pipet. Transfer the DMC fluid to a sterile 15-ml conical tube. Alternatively, DMC contents can be removed using a syringe 18-G, 1 in. needle attached to a sterile 10-ml syringe. However, aspiration of leptospires by syringe may disrupt cells and/or cause cell lysis.
    7. Examine a small aliquot of the contents DMC fluid from each chamber under dark field microscopy.
    8. Proceed to downstream applications (e.g., RNA isolation, analysis by 1D and 2D SDS-PAGE).

Representative data

Figure 3. Virulent leptospires become mammalian host-adapted during growth within dialysis membrane chambers. Representative whole cell lysates of leptospires cultivated to late-logarithmic phase in EMJH medium at 30 °C in vitro (IV) and within dialysis membrane chambers (DMC) implanted into the peritoneal cavities of female Sprague-Dawley rats. A. Lysates were loaded according to the numbers of leptospires (5 x 106 per lane) or total protein (5 µg per lane) and stained with SYPRO Ruby gel stain. Arrows and asterisks are used to highlight examples of polypeptides whose expression appears to be increased or decreased, respectively, within DMCs compared to in vitro. Molecular mass markers are indicated on the left. B. Immunoblot analyses using rabbit polyclonal antisera directed against Sph2 (Matsunaga et al., 2007), LipL32 (Haake et al., 2000) and LipL41 (Shang et al., 1996). An arrow is used to indicate a band of the predicted molecular mass for SphH, a second, closely related sphingomyelinase in L. interrogans recognized by antiserum directed against Sph2 (Matsunaga et al., 2007; Carvalho et al., 2010). Image reproduced from Caimano et al. (2014).


  1. Ketamine/xylazine anesthetic cocktail
    Stock solutions
    100 mg/ml ketamine-HCl
    20 mg/ml xylazine-HCl
  2. EMJH + BSA medium
    Prepare 1-L of EMJH medium as described in Supplementary data and previously published (Zuerner, 2005)
    Weight out 10 mg of BSA using an analytical balance
    Slowly stir in 10 mg of BSA with constant stirring, avoiding bubbles, until the BSA is completely dissolved
    Sterilize by filtration using 0.22 µm Stericup® Filter Unit
    Stored at 4 °C


The authors would like to thank Ms. Anna Allard for her superb technical assistance and Dr. Justin Radolf his continued support of our work. This work is supported by NIH Grants AI029735 (MJC), Brazilian CNPq grants 481133/2011-9 and 483052/2012-4 (AJAM) and CNPq/SWE/CSF grant 246433/2012-4 (AAG).


  1. Akins, D. R., Bourell, K. W., Caimano, M. J., Norgard, M. V. and Radolf, J. D. (1998). A new animal model for studying Lyme disease spirochetes in a mammalian host-adapted state. J Clin Invest 101(10): 2240-2250.
  2. Athanazio, D. A., Silva, E. F., Santos, C. S., Rocha, G. M., Vannier-Santos, M. A., McBride, A. J., Ko, A. I. and Reis, M. G. (2008). Rattus norvegicus as a model for persistent renal colonization by pathogenic Leptospira interrogans. Acta Trop 105(2): 176-180.
  3. Bharti, A. R., Nally, J. E., Ricaldi, J. N., Matthias, M. A., Diaz, M. M., Lovett, M. A., Levett, P. N., Gilman, R. H., Willig, M. R., Gotuzzo, E., Vinetz, J. M. and Peru-United States Leptospirosis, C. (2003). Leptospirosis: a zoonotic disease of global importance. Lancet Infect Dis 3(12): 757-771.
  4. Caimano, M. J. (2005). Cultivation of Borrelia burgdorferi in dialysis membrane chambers in rat peritonea. Curr Protoc Microbiol Chapter 12: Unit 12C 13.
  5. Caimano, M. J., Sivasankaran, S. K., Allard, A., Hurley, D., Hokamp, K., Grassmann, A. A., Hinton, J. C. and Nally, J. E. (2014). A model system for studying the transcriptomic and physiological changes associated with mammalian host-adaptation by Leptospira interrogans serovar Copenhageni. PLoS Pathog 10(3): e1004004.
  6. Carvalho, E., Barbosa, A. S., Gomez, R. M., Oliveira, M. L., Romero, E. C., Goncales, A. P., Morais, Z. M., Vasconcellos, S. A. and Ho, P. L. (2010). Evaluation of the expression and protective potential of Leptospiral sphingomyelinases. Curr Microbiol 60(2): 134-142.
  7. Haake, D. A., Chao, G., Zuerner, R. L., Barnett, J. K., Barnett, D., Mazel, M., Matsunaga, J., Levett, P. N. and Bolin, C. A. (2000). The leptospiral major outer membrane protein LipL32 is a lipoprotein expressed during mammalian infection. Infect Immun 68(4): 2276-2285.
  8. Ko, A. I., Goarant, C. and Picardeau, M. (2009). Leptospira: the dawn of the molecular genetics era for an emerging zoonotic pathogen. Nat Rev Microbiol 7(10): 736-747.
  9. Matsunaga, J., Medeiros, M. A., Sanchez, Y., Werneid, K. F. and Ko, A. I. (2007). Osmotic regulation of expression of two extracellular matrix-binding proteins and a haemolysin of Leptospira interrogans: differential effects on LigA and Sph2 extracellular release. Microbiology 153(Pt 10): 3390-3398.
  10. Shang, E. S., Summers, T. A. and Haake, D. A. (1996). Molecular cloning and sequence analysis of the gene encoding LipL41, a surface-exposed lipoprotein of pathogenic Leptospira species. Infect Immun 64(6): 2322-2330.
  11. Zuerner, R. L. (2005). Laboratory maintenance of pathogenic Leptospira. Curr Protoc Microbiol Chapter 12: Unit 12E 11.


钩端螺旋体可以感染大量的哺乳动物宿主,包括人类(Bharti等人,2003; Ko等人,2009)。在通过合适的宿主获得后,钩端螺旋体通过血流传播到多个组织,包括肾脏,在那里它们粘附并定殖在近端的回旋肾小管上(Athanazio等人。 ,2008)。感染的宿主在其尿液中排出大量螺旋体,并且螺旋体可以在传播到另一个宿主之前在不同的环境条件下存活。钩端螺旋体的差异基因表达。允许适应这些新的条件。在这里,我们描述了植入到Sprague-Dawley大鼠的腹膜腔中的透析膜室(DMC)内培养弯曲钩端螺旋体的方案(Caimano等人,2014) 。该技术最初被开发用于研究由莱姆病螺旋体,伯氏疏螺旋体(Embrynia burgdorferi)(Akins等人,1998; Caimano,2005)研究哺乳动物适应。用于该过程的透析膜管的小孔径(8,000MWCO)允许获得宿主营养物,但排除宿主抗体和免疫效应细胞。考虑到DMC和近端回旋肾小管之间的生理和环境相似性,我们推断DMC模型将适合于通过L研究体内基因表达。 interrogans 。在20至30分钟的程序中,将含有毒性钩端螺旋体的DMC手术植入大鼠腹膜腔。植入后九至11天,移出DMC并回收生物体。通常,单个DMC产生约10个哺乳动物宿主适应的钩端螺旋体(Caimano等人,2014)。除了提供用于研究病原体转录和生理变化的简易系统。询问在哺乳动物内经历,DMC模型还为选择用于诱变的新靶标和鉴定新的毒性决定簇提供了合理依据。

关键字:螺旋体, 钩端螺旋体病, 哺乳动物宿主适应性, 基因的表达, 毒力


  1. 成年雌性Sprague-Dawley大鼠(175-200g)
  2. EMJH培养基(按照补充数据中所述制备)
  3. 牛血清白蛋白(BSA)
    EMJH + BSA(10mg/ml BSA,BSA浓度20mg/ml)(Millipore,目录号:810037)
  4. 在标准体外条件(28-30 ° C)(Zuerner,2005)
  5. 超纯水(去离子,蒸馏)(例如,Milli-Q)
  6. 1mM EDTA(pH8.0)
  7. 将再生纤维素透析膜管(Spectrum Laboratories,目录号:132655)(频谱分析/目录号:132655) >
  8. 过滤单元(0.22μm,250ml,无菌)(例如,Millipore Stericup 过滤单元)
  9. 氯胺酮/甲苯噻嗪麻醉鸡尾酒(40-80mg/kg/5-10mg/kg,肌肉内给药)
  10. 眼用软膏(例如,Puralube)
  11. Betadine ®手术擦洗溶液
  12. 卡洛芬(5-10mg/kg,皮下给药)
  13. 乙醇(70%)
  14. 氯胺酮/甲苯噻嗪麻醉鸡尾酒(见配方)
  15. EMJH + BSA培养基(参见配方)


  1. 手术手套(单独包装,无菌)
  2. 一次性血清移液管(10ml无菌,单独包装)
  3. 手术巾(切成12英寸(46厘米)的方块,无菌)
  4. 纱布(4×4无菌)
  5. 配备有暗场聚光镜(例如Olympus,型号:BX40)的显微镜
  6. 三个2-L Pyrex烧杯(每个包含磁力搅拌棒)
  7. 带搅拌选项的热板
  8. 超长钝端钳(无菌)
  9. 吸管填充物(,例如,Drummond Scientific,型号:Pipet-Aid)
  10. 手术器械包,每只动物一只,使用前灭菌,并保存在无菌包装内:
    组织钳,5½英寸(〜14厘米)(1x 2齿解剖或Adson-Brown)
    组织钳,5 1/2英寸(〜14厘米)(钝端)
    带有内置剪刀(例如,Olsen Hegar)的针夹钳(5½英寸)
    缝合,Ethicon 40,SH 1,27(〜68.6厘米)涂布Vicryl,紫色编织缝线
    Autoclip ® 9毫米不锈钢伤口闭合夹和施药器
  11. 生物安全柜(生物安全2级,型号:BSL2)
  12. 循环温水毯和泵
  13. 电动毛发/毛发剪刀
  14. 玻璃珠灭菌器(例如,Braintree Scientific,型号:Germinator 500)(可选)


  1. 无菌透析膜管的制备
    1. 按照以下步骤填充三个2升Pyrex烧杯:
      烧杯1:1 - 1.5 L超纯水
      烧杯2:1-1.5L 1mM EDTA(pH 8.0)
      烧杯3:1 - 1.5 L超纯水
    2. 向每个烧杯中加入磁力搅拌棒,盖上重型铝箔和高压釜。
    3. 将每个烧杯放在热板上,在恒定搅拌下使其沸腾。
    4. 戴无菌手套,将透析膜管切成条7-9 英寸(〜18-23cm)长度使用无菌剪刀。 每切一条 动物加1-2额外。
    5. 轻轻地系住管道的一端   通过在DMC的顶部形成环而形成简单的反手结 使管的自由端通过环。 修剪掉多余的 使用无菌剪刀从绑扎端取出管子
    6. 放置绑 管插入烧杯1并更换铝箔盖。 务必 保持箔盖足够宽以允许蒸汽逸出。 煮沸管 20分钟。 不断搅拌。
    7. 使用无菌超长镊子,传输管到烧杯2.更换 铝箔盖和煮沸管20分钟。 持续搅拌。
    8. 使用烧杯3重复步骤A7。
    9. 一旦冷却到触摸,转移烧杯3到生物安全 机柜(BSC)。 为了确保无菌,含有管的烧杯 应保留在BSC中。
    注意:透析膜管可以提前几天制备并在4℃下储存。 煮沸后,使用无菌钝头镊子将透析膜管传送到0.22μmStericup过滤单元(Millipore)的底部。 在约100-200 ml的Ultrapure-Q水中更换过滤器单元顶部和过滤器。使用制造商提供的无菌帽密封过滤器单元的底部。 容器应仅在BSC内打开。

  2. 钩线虫的制备
    1. 使用标准低通量的体外毒力培养物。 interrogans),在EMJH培养基中保持在28-30℃,之后到达 在生长曲线中的稳定期
    2. 在开始外科手术之前,通过暗视野显微镜计数细菌
    3. 使用新鲜的EMJH培养基将培养物稀释至10 6个有机体/ml。
    4. 在BSC中工作,将10ml的EMJH + BSA转移到无菌的50-ml锥形管中。 每个DMC准备一个锥形管。
      注意:   重要的是补充用于DMC的EMJH培养基   额外的BSA保持适当的渗透压。
    5. 将0.1ml稀释培养物加入到含有EMJH + BSA培养基(终浓度,10μL/ml)的每个50-ml锥形管中。

  3. DMC的制备
    1. 在BSC工作,放置几个一次性,单独包装 血清学移液管在手术领域。 放下一个无菌悬垂 用作工作区。 使用自动移液器清洁表面 70%乙醇并置于无菌悬垂物上。 后穿了一双新的 无菌手术手套,松开一个50毫升锥形管的帽。
    2. 当仍然戴上手套时,小心地从其中移除一条管 储存容器使用钝端钳。 用你的自由手举行 食指和拇指之间的管道。 用其他手,删除 帽从含有稀释的细菌和转移的50-ml锥形管   使用10ml一次性血清移液管向管中加入高达9ml。   在继续保持开口端的同时,压平未填充的管 在你的手的手指之间消除任何气泡。
    3. 轻轻扭曲管道的顶端(1-2圈)以除去任何 剩余空隙体积和关闭DMC的顶部使用你的拇指 和食指握住扭曲区域以防止液体 当你打开顶部时,从DMC中泄漏出来。 绑在开放端 使用简单的反手结(例如,在DMC的顶部形成循环, 使管的自由端通过环,然后,没有 松开你的拇指和食指,轻轻地拧紧结)。 修剪 用无菌剪刀将多余的管子从绑扎端取下 注意: 尝试使DMC尽可能拉紧; 这将有助于机动 在手术期间进入腹膜腔。 结果填满 DMC应为〜1.5 - 2 in。较大的DMC可能会干扰正常 肠或膀胱功能。
    4. 将填充的DMC置于含有〜5-7ml新鲜的EMJH + BSA培养基的无菌50-ml锥形管中。
    5. 重复步骤C3-4,直到所有袋子都填满,然后直接进行手术。
      注意:   不建议储存填充的透析膜室 延长的时间,因为这可能增加的风险 污染。

  4. 腹膜植入程序。
    1. 通过肌内注射氯胺酮/赛拉嗪混合物麻醉动物
    2. 对每只眼睛涂少量眼膏。
    3. 管理术前止痛(例如,Carprofen)。
    4. 剃刮腹部并准备手术部位(例如,用Betadine外科溶液,随后用酒精连续洗涤)。
    5. 用无菌纱布覆盖手术部位,并将动物以仰卧位置转移到BSC
    6. 在开始手术前,取出覆盖的无菌纱布   手术部位和执行"脚趾捏",以确保动物是 适当麻醉。
    7. 使用无菌手术刀刀片,做一个5厘米   切口只通过皮肤,开始〜2.5厘米以下的胸腔, 使用剑突过程作为指南。 使用组织钳,拉起 皮肤在切口的任一侧并轻轻地修剪筋膜 使用解剖刀将皮肤连接到腹壁。 重复上   另一侧
    8. 使用相同的手术刀,做一个小切口(〜4厘米) 在腹壁,使用 alba 作为指南。 切口 应该干净,直。
    9. 使用组织钳,抬起一边   的腹部切口并将DMC置于大鼠腹膜腔内 (图1)。 轻轻地将DMC推向腹膜的任一侧 以防止其在缝合期间被切刻/破裂。 确定   以便其不会缠结在肠中或 干扰膀胱扩张。


    1. 通过缝合闭合腹部切口部位。 从缝合开始 每端切口有双结。 向上工作,关闭 通过放置缝合约2-3毫米留下切口
    2. 关上 皮肤切开部位使用连续的AutoClip ®伤口夹 (图2)。 注意:或者,可以关闭皮肤切除部位 使用表面下的缝合,然后是液体皮肤粘合剂(例如, Nexaband)。


    1. 将大鼠放在干净的手术巾的顶部,在一个干净的笼子里 含新鲜床上用品。 将笼子放在循环水的顶部 加热垫保持适当的核心体温。 监控 动物不断地直到警觉和反应。
    2. 止痛应该在手术后至少两天给药。
    3. 在第一周应至少每天监测动物一次 术后,然后每隔一天。 如果动物显示 任何痛苦或不适的迹象,咨询机构兽医 工作人员马上。

  5. 从DMC中恢复哺乳动物宿主适应性生物体
    1. 在植入后9至12天,通过CO 2窒息对动物实施安乐死。
    2. 将动物放在BSC卧位。
    3. 使用无菌手术剪刀,暴露腹壁。
    4. 使用无菌组织钳,抬起缝合的腹部切口部位的一侧,并使用无菌剪刀重新打开。
    5. 使用钝端钳定位和取出DMC。 转移到无菌的50毫升锥形管。
    6. 使用无菌镊子保持DMC一个结,做一个小切 在管子正下方结。 慢慢删除DMC的内容   使用无菌一次性血清移液管。 将DMC流体转移到   无菌15-ml锥形管。 或者,DMC内容可以是 使用注射器18-G,1英寸针头连接到无菌的10ml 注射器。 然而,通过注射器吸入钩端螺旋体可能破坏细胞   和/或引起细胞裂解。
    7. 在暗视野显微镜检查下,从每个腔室检查一小部分内容物DMC流体
    8. 进行下游应用(例如,RNA分离,通过1D和2D SDS-PAGE分析)。


图3.在透析膜腔室内生长期间,毒性钩端螺旋体成为哺乳动物宿主适应的。在30℃下在EMJH培养基中在体外(IV)和在植入雌性Sprague-Dawley大鼠的腹膜腔中的透析膜室(DMC)内在EMJH培养基中培养至晚对数期的螺旋体的代表性全细胞裂解物。 A.根据螺旋体数目(每个泳道5×10 6个)或总蛋白质(每个泳道5μg)加载裂解物并用SYPRO Ruby凝胶染色剂染色。箭头和星号用于突出多肽的实例,所述多肽的表达在体外与DMC相比在体外显示分别增加或减少。分子量标记在左侧显示。 B.使用针对Sph2的兔多克隆抗血清(Matsunaga等人,2007),LipL32(Haake等人,2000)和LipL41(上海)的免疫印迹分析, et al。,1996)。箭头用于指示SphH的预测分子量的带,第二,密切相关的鞘磷脂酶在L中。 (Matsunaga等人,2007; Carvalho等人,2010)通过针对Sph2的抗血清识别的抗原特异性抗体来自Caimano等人的图片(2014)。


  1. 氯胺酮/甲苯噻嗪麻醉剂
    100mg/ml氯胺酮-HCl 20mg/ml甲苯噻嗪-HCl
  2. EMJH + BSA培养基
    减去10mg BSA 在持续搅拌下在10mg BSA中缓慢搅拌,避免气泡,直到BSA完全溶解为止 使用0.22μmStericup 过滤单元
    过滤灭菌 储存在4°C


作者感谢Anna Allard女士的杰出技术援助,并感谢Justin Radolf博士继续支持我们的工作。 这项工作由NIH Grants AI029735(MJC),巴西CNPq拨款481133/2011-9和483052/2012-4(AJAM)和CNPq/SWE/CSF拨款246433/2012-4(AAG)支持。


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引用:Grassmann, A. A., McBride, A. J., Nally, J. E. and Caimano, M. J. (2015). Generation of Mammalian Host-adapted Leptospira interrogans by Cultivation in Peritoneal Dialysis Membrane Chamber Implantation in Rats. Bio-protocol 5(14): e1536. DOI: 10.21769/BioProtoc.1536.