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Sebinger Culture: A System Optimized for Morphological Maturation and Imaging of Cultured Mouse Metanephric Primordia
Sebinger培养:一个用于小鼠后肾原基形态成熟和成像的优化系统   

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PLOS ONE
May 2010

Abstract

Here, we present a detailed protocol on setting up embryonic renal organ cultures using a culture method that we have optimised for anatomical maturation and imaging. Our culture method places kidney rudiments on glass in a thin film of medium, which results in very flat cultures with all tubules in the same image plane. For reasons not yet understood, this technique results in improved renal maturation compared to traditional techniques. Typically, this protocol will result in an organ formed with distinct cortical and medullary regions as well as elongated, correctly positioned loops of Henle. This article describes our method and provides detailed advice. We have published qualitative and quantitative evaluations on the performance of the technique in Sebinger et al. (2010) and Chang and Davies (2012).

Keywords: Organ culture (器官培养), Kidney (肾), Metanephros (后肾), Sebinger culture (Sebinger培养), Organoid (类器官), Imaging (成像)

Background

The metanephric (permanent) kidneys of mammals develop from simple rudiments located at the caudal end of the intermediate mesoderm. In mice, at about embryonic day (‘E’) 10 these rudiments form and consist of two morphologically distinguishable components; an epithelial ureteric bud that arises as a diverticulum of the Wolffian (nephric) duct, and a metanephrogenic mesenchyme that forms next to the duct. As development progresses, the ureteric bud enters the metanephrogenic mesenchyme and undergoes many successive rounds of growth and branching to make a ‘tree’; this later remodels to produce a mature collecting duct system in which tubules radiate from a central cavity, the renal pelvis (Lindstrom et al., 2015). The renal pelvis drains to the ureter, which forms from the original stalk of the ureteric bud. As the ureteric bud develops, it induces cells from the metanephrogenic mesenchyme to condense around each of its tips to form a ‘cap mesenchyme’ (Schreiner, 1902; Reinhoff, 1922). The cap mesenchymes are stem cell populations that divide as the tips divide, so that each tip formed by bifurcation of an existing branch inherits its own cap (reviewed by Hendry et al., 2011). Cells at the more distal ends of the caps differentiate to form excretory nephrons, which connect to the ureteric bud branch from whose cap they formed (Georgas et al., 2009). Blood vessels invade from the base of the metanephros and follow the ureteric bud, making a network around (but never entering) the cap mesenchymes (Munro et al., 2017): later, these vessels will serve glomeruli and other parts of the kidney.

In vitro culture of metanephric kidney rudiments has a long history. Indeed, these were among the first embryonic organs to show continued development outside the body (Carrel and Burrows, 1910). The continued development of kidney rudiments outside of the body was a scientific, as well as a technical, advance: the autonomous development of isolated organs demonstrated that the information required to build them was ‘local’ and did not depend on the rest of the embryo. This observation added considerable support to the idea that architecture of the body is hierarchical, with modules (organs) that largely look after themselves and interact with the rest of the body only at specific functional interfaces. The earliest culture methods used rather complex media and culture supports, such as Grobstein’s use of clotted avian plasma and chick embryo extract (Grobstein, 1953). These systems were necessary because simply placing a kidney rudiment in a glass dish or flask resulted in its breakdown because cells adhered to the substrate and spread out to form a monolayer. Immersion in medium in non-adhesive dishes prevents cell dispersal but does not result in proper development (see ‘Data analysis’ section). Both imaging and reproducibility were greatly improved by Saxén’s adoption of a culture method developed by Trowell for culture of rat lymph nodes (Trowell, 1954). Saxén placed embryonic kidney rudiments, isolated from mice at E11, on filters that were supported by a stainless steel grid at the interface between gas and medium (Saxén et al., 1962).

The Trowell method has been a mainstay of research in kidney development for many decades. It allows for significant ureteric bud growth and branching, formation of nephrons, differentiation of their separate proximal and distal domains and connection of nephrons to ureteric bud branches. Rudiments grow flat enough to facilitate confocal microscopy of fixed specimens without the need for ‘clearing’ techniques, and even conventional epifluorescence microscopy is adequate for many purposes (e.g., Davies et al., 2014). Another advantage is that diffusion paths have proved to be short enough and open enough to enable mechanisms of development to be investigated in this method with drugs (e.g., Fisher et al., 2001), growth factors (e.g., Piscione et al., 1997), function-blocking antibodies (e.g., Falk et al., 1996) and, to a limited extent, siRNAs (e.g., Davies et al., 2004). The Trowell system, together with variants that place rudiments at the air-medium interface using use Transwell filters instead of stainless steel supports, remains very common in studies of kidney development.

Useful as it is, the Trowell culture system has a few problems. One is that the filter itself interferes with bright-field/phase contrast imaging because the filter pores appear, out of focus, in the image (though some Transwell systems do achieve good optical clarity). Another is that the tissue is too thick for reliable high-resolution, unattended time-lapse photography, because tubules leave the focal plane. A third problem is that some aspects of renal development, such as the formation of distinct cortex and medulla, and extension of nephrons’ loops of Henle from the cortex to the medulla, occur poorly if at all. In an attempt to address these issues, we developed an alternative culture system that uses the surface tension of very shallow medium to hold a kidney rudiment on to a clear glass substrate. To our surprise, the system not only solved the imaging problem, but it also allowed the organ rudiments to develop clear cortico-medullary zonation and nephron maturation proceeded as far as the production of clear and elongated loops of Henle (Sebinger et al., 2010) (Figure 1). The enhanced development is seen in cultures made from natural kidneys isolated from mouse embryos, and also in organoids engineered from suspensions of stem cells (Chang and Davies, 2012).


Figure 1. Kidneys cultured in the Sebinger system. A. Bright field images for E11.5 kidney grown in culture for 0, 3 and 7 days. Scale bars = 0.5 mm. B-D. E11.5 kidneys cultured for 7 days in the Sebinger system and stained for different renal markers to show maturation. B. Stained for the ureteric bud marker CALB (shown in green) and the basement membrane marker Laminin (shown in red). The red channel shows the presence of loops of Henle dipping into the medulla; C. Stained for CALB (green) and the proximal tubular marker LTL (red); D. Stained for ECAD (ureteric bud and distal tubular marker; shown in green) and WT1 (podocyte and cap mesenchyme marker; shown in red). Scale bars = 100 μm.

Materials and Reagents

  1. 1 ml disposable syringes (Plastipak 1 ml, BD, catalog number: 303172 ) with fine needles (0.5 x 16 mm/25 G x 5/8”, Microlance 3, BD, catalog number: 300600 )
  2. 40 x 0.13 mm borosilicate glass coverslips (VWR, catalog number: 631-0177 )
  3. Silicone cones (flexiPERM Cone shape A, Greiner Bio One International)
    These are at the time of writing available only on special order–phone Greiner–and delivery times can be long enough to make forward planning important. The cones can be re-used for years.
    Note: We know of no suitable substitutes.
  4. 100 mm sterile Petri dishes (for dissection–surface quality is irrelevant) (Cell Star®, Greiner Bio One International, catalog number: 664160 )
  5. 60 x 15 mm sterile Petri dishes (Cell Star®, Greiner Bio One International, catalog number: 628160 )
    Note: We use Greiner but expect that others will also be suitable.
  6. Glass Pasteur pipettes (150 mm, Volac, catalog number: D810 )
  7. Timed-mated mice at E11.5 (E10.5 is also suitable but is a more challenging dissection)
  8. Sterile distilled water
  9. 100% methanol
  10. Eagle’s minimal essential medium with Earle’s salts and non-essential amino acids (Sigma-Aldrich, catalog number: M5650 )
  11. Foetal bovine serum (FBS) (Biochrom, catalog number: S 0415 )
  12. Penicillin-streptomycin (Sigma-Aldrich, catalog number: P4333 )
  13. Phosphate buffered saline (PBS) (Sigma-Aldrich, catalog number: 79382 )
  14. Anti-laminin (Sigma-Aldrich, catalog number: L9393 )
  15. FITC anti-rabbit (Sigma-Aldrich, catalog number: F0382 )
  16. EmbryoMax® Penicillin-streptomycin solution, 100x (Sigma-Aldrich, catalog number: TMS-AB2-C )
  17. Hydrogen peroxide (Sigma-Aldrich, catalog number: H1009 )
  18. Ammonium hydroxide (Sigma-Aldrich, catalog number: A6899 )
  19. Cleaning solution (see Recipes)
  20. Dissecting medium (see Recipes)
  21. Culture medium (see Recipes)
  22. Hydration buffer (see Recipes)

Equipment

  1. Forceps
  2. Scalpel (curved blade, e.g., D-form, type 22, Swann Morton, catalog number: 0108 )
  3. 80 °C oven
    Note: We use a Gallenkamp Hotbox size 1, but any 80 °C oven should be suitable.
  4. Cell culture incubator at 37 °C, 5% CO2
    Note: We use a NuAire 5500E (NuAire, model: NU-5500E ), but any stable incubator should work as well.
  5. Dissecting microscopes
    Note: We use ZEISS Stemi 2000C microscopes (ZEISS, model: Stemi 2000-C ) with transilluminating stages, but have demonstrated the technique in other laboratories with other models of dissecting microscope. The precise type of microscope is not important, but transillumination (rather than epi-illumination) is.
  6. Clean area
    Note: We do not use safety cabinets (for the non-pathogen-infected samples we use), because the vibration of the fans is a nuisance. We do use simple cabinets about the size of a safety cabinet, made from Perspex, to provide shelter from dust moved by Edinburgh winds blowing through ill-fitting antique lab windows).

Procedure

  1. Preparation
    1. Make the solutions listed in Recipes section.
    2. Sterilize 40 x 0.13 mm coverslips (depending on the manufacturer, they may need to be acid treated in 1 N HCl first; 10 min room temperature, followed by 3 rinses in sterile distilled water).
    3. Clean the flexiPERM cones in cleaning solution (50 ml or so–the exact volume does not matter) at 70 °C for 10 min. Rinse them 3 x and store them in sterile distilled water.

  2. Isolation of E11.5 mouse kidney rudiments
    Note: Takes about 20 min per pregnant mouse in skilled hands, a lot longer for beginners. Absolute beginners are advised to begin by isolating E12.5 kidneys before moving on to harder-to-see E11.5 kidneys. All steps should be done under sterile conditions.
    1. Sacrifice the pregnant mother mouse by methods appropriate to the local animal licence and laws.
    2. Remove the uterus.
    3. In a large Petri dish, immerse the uteri in dissecting medium and use a scalpel to cut between bulges in the uterus: ‘rolling’ the curved edge of the scalpel over the tissue to be cut is easier than pulling it in the conventional way. Squeeze the embryo out of the cut end next to it with the forceps. Remove the heads of the embryos, and any placental/membrane material. (Figure 2)


      Figure 2. Illustration of the E11.5 metanephric kidney isolation. A. The pregnant uterus; B. Squeezing embryos out of the uterus with forceps; C. The extracted E11.5 embryos; D. The caudal part of the embryo; E. The rear of the embryo, after cutting it sagittally into two halves; the arrow points towards the metanephric kidney, edges of which are marked in white in the detail (seeing the kidney is the hardest part of the dissection, and takes practise). F. The isolated E11.5 metanephric kidney. Scale bar for A-C is 5 mm, for D and E is 1 mm, and 0.5 mm for F.

    4. Transfer the embryo trunks to a 60 mm dish filled to about 5 mm depth with dissecting medium (the precise amount can be altered to suit individual preferences), which can be used straight from storage at 4 °C.
    5. Using fine needles as dissecting instruments (see Materials and Reagents for details), transect the embryos just rostral to the hind limb buds. Retain the portions with the hind limb buds (these can be stored for a few days or transported on ice: see Davies, 2006, for details of the method).
    6. Remove the tails (which are otherwise a springy nuisance while dissecting).
    7. Lay each rear-end fragment of the embryo ventral-side-down, and cut it sagittally, directly down from the mid-line of the neural tube to its ventral surface.
    8. Inspect each half created in the last step, from the cut (anatomically medial) surface, and identify the kidney rudiment, which lies (at E11.5) next to the cranial limit of the hind limb bud. The ureteric bud diverticulum of the nephric duct is the most obvious spatial cue, and the extend of the metanephrogenic mesenchyme can be seen by a slight change in light-scattering properties (on some microscopes it appears lighter than the surrounding mesoderm, and on some it appears darker).
    9. Dissect the kidney rudiment free of the rest of the embryo, pinning tissue down with one needle and drawing the other needle against it to make accurate cuts, and transfer the kidney to a new dish in which rudiments can be pooled for later use (pulled glass pipettes provide one means of transferring rudiments easily: the heat of melting the glass to pull them will sterilize their ends).
      Note: Throughout the dissection, care should be taken to ensure that media are not left in air for so long that they become alkaline (the pH indicator in the medium will indicate this by a colour change. We suggest 15 min as a maximum but air movement may mean shorter times are needed. In our experience, HEPES inhibits normal kidney development and we avoid its use).

  3. Sebinger culture
    1. Put the cover slips and the silicone cones in 80 °C oven for 20 min to dry (dryness is essential for proper adhesion between the silicone cone and the cover slips). They can be handled with gloved hands or forceps–use plastic forceps for cones to avoid damage that might result from the use of metal.
    2. Place a cone, narrow-end down, on to the coverslip and use a blunt-ended forceps to press the edge of the silicone cone against the cover slip to ensure tight adhesion.
    3. Place the cover slip with the silicone cone attached to it on the base of a 60 mm Petri dish. (Figure 3)


      Figure 3. The Sebinger culture method. A. Shows the different component of the culture system. B. Shows the assembled Sebinger culture system.

    4. Pipette a kidney rudiment into the approximate centre of the glass circle defined by the cone.
    5. Working quickly to avoid the risk of the tissue drying out, pipette away any medium carried over with the kidney rudiment, and place 85 μl of culture medium in the circle. It can be used straight from cold storage. Use a pipette tip to ensure that this medium spreads to cover the whole of the circle of glass bounded by the cone, including the kidney, and does not ‘bead’ into a single drop (Figure 4). Note that the optimal amount of medium varies slightly with batches of cones and with the glass. We therefore recommend that users try a series of volumes (e.g., 80, 82.5, 85. 97.5, 90 μl) with their own materials to determine the optimum.
    6. Place the 60 mm dish (with the cover slip, the silicone cone and the kidney rudiment now inside it) to the base of a 100 mm Petri dish and add 3 ml of hydration buffer into it (when the hydration buffer comes in contact with the silicone cone it detaches the cone from the cover slip and the system becomes leaky, this is why we use a separate dish to add the hydration buffer). (Figure 4)


      Figure 4. The arrangement of the cone and dishes. A. Kidney rudiments are cultured in the centre of the silicone cone in a low volume of KCM medium and a humidifying buffer is added to the outer dish to prevent dryness. B. The KCM tends to bead as a single drop (left hand side image) and need to be distributed with a pipette tip to cover the glass circle enclosed by the silicone cone (right hand side image).

    7. Place the lid on the 100 mm Petri dish then place the dish in a tissue culture incubator (37 °C, 5% CO2, 100% humidity). Kidneys can be cultured for 3 days; beyond that it is advisable to change the medium. A few cells at the periphery may egress and make a monolayer on the glass but the kidney as a whole should remain intact and coherent, as in Figure 1A. If it ‘rounds up’ but is still alive there is probably too much medium; if it rounds up as a dried mass, there is too little.
      Note: For time-lapse filming, we recommend in-incubator microscopes from Etaluma.
    8. Specimens should be fixed on their cover-slip, according to the protocol appropriate for any antibody staining to be used. We generally apply 100% methanol at -20 °C immediately after removing the culture medium, and allow it to warm to room temperature over 15 min, then wash in phosphate-buffered saline. If alternative fixation techniques involve detergents (Tween, Triton etc.), great care should be taken to change solutions very gently to avoid detaching the culture from the glass. Incubation in antibody is usually overnight at 4 °C in Bijou (5 ml) bottles. An example protocol, used for the laminin stain in Figure 1, is overnight incubation in 200 μl 1/100 anti-laminin (Sigma-Aldrich) in PBS at 4 °C, a 7 h wash in PBS at 4 °C, overnight incubation in 200 μl 1/100 FITC anti-rabbit (Sigma-Aldrich) in PBS at 4 °C, two 2 h washes in PBS at 4 °C, followed by mounting, still on the filter, on a slide in PBS or 1:1 PBS-glycerol, with the cover-slip being held away from the main slide with fragments a broken coverslip used as spacers. This prevents crushing of the tissue. We do not embed or section.

Data analysis

  1. This is a method of culture, rather than a method to make a specific measurement, and the method will support a range of questions and analyses. Its features do, however, make it better suited for some questions than others. For avoidance of false-positive data caused by individual variation, if embryos from more than one mother mouse are to be used we recommend pooling kidneys from all the embryos, followed by random allocation to experimental and control groups.
  2. The extent of manual manipulation involved in the method makes the final size and shape of the organ rudiment quite variable, and some do not grow at all, usually because the film of medium has become a drop as in Figure 4B, left panel. At the very least, experimenters need to settle some exclusion criteria so that failed cultures can be excluded fairly from analyses (ideally, before any analyses are made except for simple observation of the medium film). Variation in growth between samples, probably arising from tiny differences in placement of specimens in the film, meaning that measurements of features such as total organ area will generate large error bars and require many replicates before any meaningful comparisons of experimental and control kidneys can be made. We advise against using such a measure. Better quantitative measurements include the number of branch tips in the collecting duct tree, and the number of nephrons. Examples of branch tip counting analysis can be found in Fisher et al. (2001) and Michael et al. (2005). Even for this, there is always an ambiguity about how ‘T-shaped’ a branch end must be for it to be counted as two tips not one, and there is always some ambiguity about what is mature enough to be called a nephron. We therefore strongly advise that all samples are blind-coded or, if this is not practical, images of them are blind-coded and counted by someone who does not know whether samples come from experiment or control.
  3. Much information gained from observation of organ culture, whether normal or subject to experimental manipulation, is qualitative rather than quantitative: tubular morphology is an example. Again, researchers are encouraged to use blind-coding, perhaps using images viewed by multiple independent people, and perhaps to use a semi-quantitative scoring system even for morphology (for example, classification of a nephron as ‘normal’ or ‘abnormal’). Some data are about directions, for example the direction in which a Loop of Henle grows. Directions require a coordinate system. A simple coordinate system can be made by extending lines radially from, for example, the centre of the kidney (determined, for example, using ImageJ’s Centre of Mass function), or from the first branching point. Directions of loops of Henle, for example, can be measured in terms of the angle between the tubule of the loop and the radial line passing through its tip. Again, having multiple biological replicates and multiple people making measurements on the same blind-coded samples will provide good measurements of inter-observer variability and of inter-sample variability.

Recipes

  1. Cleaning solution
    This consists of 5:1:1 (by volume) H2O:H2O2:NH4OH
    Make this by adding the hydrogen peroxide and ammonium hydroxide to the water, not the other way round, and use personal protective equipment (gloves, eye protection) when handling the concentrated solutions
  2. Dissecting medium
    Dissolve Eagle’s minimal essential medium with Earle’s salts and non-essential amino acids in ddH2O as per manufacturer’s instruction
  3. Culture medium: KCM
    Note: Simply add the serum and antibiotics to the Eagle’s medium (Dissecting medium).
    Dissecting medium
    10% fetal bovine serum
    1/100 penicillin/streptomycin stock (this stock contains 10.000 U/ml penicillin and 10 mg/ml streptomycin)
  4. Hydration buffer
    PBS (from Sigma-Aldrich tablets: allow an hour for the tablets to dissolve, and mix well, then add penicillin/streptomycin diluted from the 100x stock mentioned in Recipe 3 above)

Acknowledgments

Development and use of the technique described here has been supported by funding from the following; European Commission FP6 grant MRTN-CT-2006-036097, Medical Research Council grant MR/K010735/1, Kidney Research UK grant RP_002_20160223, and a PhD scholarship from the Egyptian Educational and Cultural Bureau. The protocol was first described in the methods section of Sebinger, 2010. The authors have no competing interests.

References

  1. Carrel, A. and Burrows, M. T. (1910). Cultivation of adult tissues and organs outside the body. J Am Med Ass 55: 1379-1381.
  2. Chang, C. H. and Davies, J. A. (2012). An improved method of renal tissue engineering, by combining renal dissociation and reaggregation with a low-volume culture technique, results in development of engineered kidneys complete with loops of Henle. Nephron Exp Nephrol 121(3-4): e79-85.
  3. Davies, J. A. (2006). A method for cold storage and transport of viable embryonic kidney rudiments. Kidney Int 70(11): 2031-2034.
  4. Davies, J. A., Hohenstein, P., Chang, C. H. and Berry, R. (2014). A self-avoidance mechanism in patterning of the urinary collecting duct tree. BMC Dev Biol 14: 35.
  5. Davies, J. A., Ladomery, M., Hohenstein, P., Michael, L., Shafe, A., Spraggon, L. and Hastie, N. (2012). Development of an siRNA-based method for repressing specific genes in renal organ culture and its use to show that the Wt1 tumour suppressor is required for nephron differentiation. Hum Mol Genet 2004: 235-46.
  6. Falk, M., Salmivirta, K., Durbeej, M., Larsson, E., Ekblom, M., Vestweber, D. and Ekblom, P. (1996). Integrin alpha 6B beta 1 is involved in kidney tubulogenesis in vitro. J Cell Sci 109 (Pt 12): 2801-2810.
  7. Fisher, C. E., Michael, L., Barnett, M. W. and Davies, J. A. (2001). Erk MAP kinase regulates branching morphogenesis in the developing mouse kidney. Development 128(21): 4329-4338.
  8. Georgas, K., Rumballe, B., Valerius, M. T., Chiu, H. S., Thiagarajan, R. D., Lesieur, E., Aronow, B. J., Brunskill, E. W., Combes, A. N., Tang, D., Taylor, D., Grimmond, S. M., Potter, S. S., McMahon, A. P. and Little, M. H. (2009). Analysis of early nephron patterning reveals a role for distal RV proliferation in fusion to the ureteric tip via a cap mesenchyme-derived connecting segment. Dev Biol 332(2): 273-286.
  9. Grobstein, C. (1953). Inductive epithelio-mesenchymal interaction in cultured organ rudiments in the mouse. Science 118: 52-55.
  10. Hendry, C., Rumballe, B., Moritz, K. and Little, M. H. (2011). Defining and redefining the nephron progenitor population. Pediatr Nephrol 26(9): 1395-1406.
  11. Lindstrom, N. O., Chang, C. H., Valerius, M. T., Hohenstein, P. and Davies, J. A. (2015). Node retraction during patterning of the urinary collecting duct system. J Anat 226(1): 13-21.
  12. Michael, L., Sweeney, D. E. and Davies, J. A. (2005). A role for microfilament-based contraction in branching morphogenesis of the ureteric bud. Kidney Int 68(5): 2010-2018.
  13. Munro, D. A. D., Hohenstein, P. and Davies, J. A. (2017). Cycles of vascular plexus formation within the nephrogenic zone of the developing mouse kidney. Sci Rep 7(1): 3273.
  14. Piscione, T. D., Yager, T. D., Gupta, I. R., Grinfeld, B., Pei, Y., Attisano, L., Wrana, J. L. and Rosenblum, N. D. (1997). BMP-2 and OP-1 exert direct and opposite effects on renal branching morphogenesis. Am J Physiol 273(6 Pt 2): F961-975.
  15. Reinhoff, W. F. (1922). Development and growth of the metanephros or permanent kidney in chick embryos. Johns Hopkins Hospital Bulletin 33: 392-406.
  16. Saxén, L., Vainio, T. and Toivonen, S. (1962). Effect of polyoma virus on mouse kidney rudiment in vitro. J Natl Cancer Inst 29: 597-631.
  17. Schreiner, K. E. (1902). Ueber die entwicklung der amniotenniere. Zeitsch. f. wiss. Zool. 71: 1-188.
  18. Sebinger, D. D., Unbekandt, M., Ganeva, V. V., Ofenbauer, A., Werner, C. and Davies, J. A. (2010). A novel, low-volume method for organ culture of embryonic kidneys that allows development of cortico-medullary anatomical organization. PLoS One 5(5): e10550.
  19. Trowell, O. A. (1954). A modified technique for organ culture in vitro. Exp Cell Res 6(1): 246-248.

简介

在这里,我们提出了一个详细的协议,建立胚胎肾脏器官培养使用培养方法,我们已经优化解剖成熟和成像。 我们的培养方法是将肾脏的基质放在玻璃上,形成一层薄薄的培养基,培养的平面非常平坦,所有的肾小管都在同一图像平面上。 由于尚未理解的原因,与传统技术相比,该技术导致肾成熟的改善。 通常情况下,这个协议将导致器官形成不同的皮层和髓质区域,以及拉长,正确定位的亨利循环。 本文介绍了我们的方法并提供了详细的建议。 我们已经在Sebinger等人(2010)和Chang和Davies(2012)上发表了关于该技术性能的定性和定量评估。

【背景】哺乳动物的后肾(永久性)肾发育于位于中胚层尾端的简单遗传。在小鼠胚胎日('E')10,这些基因形成并由两种形态上可区分的组分组成;产生为Wolffian(肾)导管的憩室的上皮性输尿管芽,以及形成在导管旁的后肾间质(metanephrogenic mesenchyme)。随着发育进展,输尿管芽进入后肾间质,经历多轮生长和分枝,形成“树”。这后来重塑以产生一个成熟的集合管系统,其中肾小管从中心腔,肾盂放射(Lindstrom等人,2015)。肾盂排水到输尿管,从输尿管芽的原始茎形成。随着输尿管芽发育,它诱导来自后肾间充质细胞的细胞在其每个尖端周围凝结形成“帽间充质”(Schreiner,1902; Reinhoff,1922)。帽状间充质是干细胞群体,随着尖端分裂而分裂,因此通过分支形成的每个尖端继承自身的帽子(由Hendry等人2011年综述)。帽的更远端的细胞分化形成排泄肾单位,这些肾单位连接到他们形成的输尿管芽分支(Georgas et al。,2009)。血管从肾后基底侵入并跟随输尿管芽,在肾间质周围形成网状结构(但从未进入)(Munro等人,2017):之后,这些血管将为肾小球和肾脏的其他部分。

后肾肾衰的体外培养有着悠久的历史。事实上,这些是第一个胚胎发育器官之间的持续发展(Carrel和Burrows,1910)。肾脏遗体在体外的持续发展是科学的,也是一个技术上的进步:孤立器官的自主发展表明,构建它们所需要的信息是“本地的”,并且不依赖胚胎的其余部分。这一观察结果为人们的体系结构是分层的观点增加了很大的支持,模块(器官)很大程度上照顾自己,只在特定的功能界面与身体其他部分相互作用。最早的培养方法使用相当复杂的培养基和培养基,例如Grobstein使用凝血禽血浆和鸡胚提取物(Grobstein,1953)。这些系统是必要的,因为简单地将肾脏基质置于玻璃皿或烧瓶中导致其分解,因为细胞粘附到基底并展开形成单层。浸泡在不粘合的培养皿中防止细胞扩散,但不会导致适当的发育(参见“数据分析”部分)。 Saxén采用Trowell培养的大鼠淋巴结培养方法(Trowell,1954)显着改善了成像和重现性。 Saxén将从E11小鼠分离出来的胚胎肾脏原始物质放置在由气体和介质之间的界面处的不锈钢格栅支撑的过滤器上(Saxén等人,1962)。

Trowell方法几十年来一直是肾脏发育研究的主流。它允许显着的输尿管芽生长和分支,肾单位的形成,他们分开的近端和远端域的分化以及肾单位与输尿管芽分支的连接。原生质体的生长足够平坦,以便于固定标本的共焦显微镜检查,而不需要“清除”技术,甚至传统的落射荧光显微镜也适用于许多目的(例如,Davies 等,2014)。另一个优点是扩散路径已经被证明足够短并且足够开放以使得可以用药物(例如,Fisher等人, 2001),生长因子(例如,Piscione等人,1997),功能阻断抗体(例如Falk等人, 。,1996),并且在一定程度上是siRNAs(例如,Davies等人,2004)。 Trowell系统以及使用Transwell过滤器代替不锈钢支撑物在空气 - 介质界面上放置雏形的变体在肾脏发育的研究中仍然是非常普遍的。

Trowell文化系统很有用,但有一些问题。一个是过滤器本身干扰明场/相衬成像,因为过滤器毛孔在图像中出现失焦(虽然一些Transwell系统确实获得了良好的光学清晰度)。另一个原因是由于小管离开焦平面,组织对于可靠的高分辨率,无人照管的延时摄影而言太厚。第三个问题是肾发育的某些方面,例如形成不同的皮层和髓质,以及Henle的肾单位从皮层延伸到延髓的延伸,如果有的话,则发生得不好。为了解决这些问题,我们开发了一种替代的培养系统,利用非常浅的培养基的表面张力将肾脏基质保持在透明的玻璃基质上。令我们惊讶的是,该系统不仅解决了成像问题,而且还允许器官基因发育明显的皮质髓质分带和肾单位成熟,直至产生Henle的清晰和延长的环(Sebinger等人。,2010)(图1)。从小鼠胚胎分离出来的天然肾脏以及由干细胞悬浮液工程化的类固醇(Chang and Davies,2012)中可以看到增强的发展。

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图1.在Sebinger系统中培养的肾脏A.培养0天,3天和7天的E11.5肾的明场图像。比例尺= 0.5毫米。 B-d。 E11.5肾在Sebinger系统中培养7天,并对不同的肾标志物进行染色以显示成熟。 B.输尿管芽标记CALB(显示为绿色)和基底膜标记层粘连蛋白(显示为红色)染色。红色的通道显示了Henle的环路浸入髓质的存在; C. CALB染色(绿色)和近端管状标记LTL(红色); D.对于ECAD(输尿管芽和远端管状标记;以绿色显示)和WT1(足细胞和帽间充质标记;以红色显示)染色。比例尺= 100微米。

关键字:器官培养, 肾, 后肾, Sebinger培养, 类器官, 成像

材料和试剂

  1. 使用细针(0.5×16mm / 25G×5/8“,Microlance 3,BD,目录号:300600)的1ml一次性注射器(Plastipak 1ml,BD,目录号:303172)
  2. 40 x 0.13 mm硼硅酸盐玻璃盖玻片(VWR,目录号:631-0177)
  3. 硅胶锥(flexiPERM锥形A,格瑞纳生物一国际)
    在撰写本文时,只能通过Greiner的特殊订购电话进行交付,并且交货时间可以足够长,以使前瞻性计划变得重要。锥体可以重复使用多年。
    注:我们知道没有合适的替代品。
  4. 100毫米无菌培养皿(用于解剖 - 表面质量无关紧要)(Cell Star ,Greiner Bio One International,目录号:664160)
  5. 60×15毫米无菌培养皿(Cell Star ,Greiner Bio One International,目录号:628160)
    注意:我们使用Greiner,但期望别人也适合。
  6. 玻璃巴斯德吸液管(150毫米,沃拉克,目录号:D810)
  7. 在E11.5(E10.5也是合适的,但更具挑战性的解剖)定时交配老鼠。
  8. 无菌蒸馏水
  9. 100%甲醇
  10. Eagle's最小基本培养基,含有Earle's盐和非必需氨基酸(Sigma-Aldrich,目录号:M5650)
  11. 胎牛血清(FBS)(Biochrom,目录号:S 0415)
  12. 青霉素 - 链霉素(Sigma-Aldrich,目录号:P4333)
  13. 磷酸盐缓冲盐水(PBS)(Sigma-Aldrich,目录号:79382)
  14. 抗层粘连蛋白(Sigma-Aldrich,目录号:L9393)
  15. FITC抗兔(Sigma-Aldrich,目录号:F0382)
  16. 胚胎最大青霉素 - 链霉素溶液,100x(Sigma-Aldrich,目录号:TMS-AB2-C)
  17. 过氧化氢(Sigma-Aldrich,目录号:H1009)
  18. 氢氧化铵(Sigma-Aldrich,目录号:A6899)
  19. 清洁解决方案(见食谱)
  20. 解剖介质(见食谱)
  21. 培养基(见食谱)
  22. 水合缓冲液(见食谱)

设备

  1. 镊子
  2. 手术刀(弯曲刀片,例如,D型,22型,Swann Morton,目录号:0108)
  3. 80°C烤箱
    注意:我们使用1号Gallenkamp Hotbox,但任何80°C的烤箱都应该是合适的。
  4. 细胞培养箱在37°C,5%CO 2
    注意:我们使用NuAire 5500E(NuAire,型号:NU-5500E),但任何稳定的培养箱都应该能够正常工作。
  5. 解剖显微镜
    注:我们使用ZEISS Stemi 2000C显微镜(ZEISS,型号:Stemi 2000-C)和透照阶段,但在其他实验室用其他解剖显微镜模型显示了该技术。显微镜的精确类型并不重要,但透照(而不是落射照明)是。
  6. 清洁区域
    注:我们不使用安全柜(对于我们使用的非病原体感染的样本),因为风扇的振动是一个麻烦。我们确实使用与Perspex制造的安全柜尺寸相似的简易橱柜,以防止爱丁堡风吹过不合身古色古香的实验室窗户的灰尘。

程序

  1. 制备
    1. 将解决方案列在“食谱”部分。
    2. 消毒40×0.13毫米的盖玻片(取决于制造商,他们可能需要首先在1N HCl中进行酸处理;室温10分钟,然后在无菌蒸馏水中漂洗3次)。
    3. 清洁溶液中的flexiPERM锥体(50毫升左右 - 确切的体积无关紧要)在70°C下10分钟。冲洗3次,并将其储存在无菌蒸馏水中。

  2. 分离E11.5小鼠肾衰竭
    注意:每个怀孕的老鼠在熟练的手中花费大约20分钟,对于初学者来说,花费的时间要长得多。建议绝对的初学者首先分离E12.5肾脏,然后再转向难以看见的E11.5肾脏。所有步骤应在无菌条件下进行。
    1. 根据当地的动物许可证和法律,牺牲怀孕的母鼠。
    2. 去除子宫。
    3. 在一个大的培养皿中,将子宫浸泡在解剖介质中,并用手术刀在子宫的凸起之间切割:在待切割的组织上“滚动”手术刀的弯曲边缘比以常规方式拉动手术更容易。用钳子将胚胎挤压出来。移除胚胎的头部,以及任何胎盘/膜材料。 (图2)


      图2.E11.5后肾肾分离图。 :一种。怀孕的子宫; B.用钳子将胚胎挤出子宫; C.提取的E11.5胚胎; D.胚胎的尾部; E.胚胎的后方,切成两半;箭头指向后肾,其边缘在细节上标记为白色(看到肾脏是解剖中最难的部分,并且需要练习)。 F.分离的E11.5后肾。 A-C的比例尺为5毫米,D和E为1毫米,F为0.5毫米。

    4. 将胚胎移植到一个直径约5毫米的60毫米培养皿中,使用解剖介质(精确的数量可以根据个人喜好进行调整),可以在4°C下直接使用。
    5. 使用细针作为解剖工具(详见材料和试剂),将胚胎横切至后肢芽。留下后肢芽的部分(这些可以存储几天或在冰上运输:见戴维斯,2006年,方法的细节)。

    6. 删除尾巴(否则这是一个松懈的问题,同时解剖)
    7. 将胚胎腹侧的每个后端碎片放下,并从神经管的中线直接向下切到腹面。
    8. 检查在最后一步创建的每一半,从切割(解剖学中度)的表面,并确定(在E11.5)在后肢芽的头界限旁边的肾脏的雏形。肾管的输尿管芽憩室是最明显的空间线索,并且可以通过光散射特性的轻微改变来看到后肾间质的延伸(在一些显微镜上,它看起来比周围中胚层更轻,并且在一些上看起来更黑)。
    9. 解剖胚胎的其余部分的肾脏,用一根针将组织固定下来,并将另一根针头靠在它上面以便精确切割,并将肾脏转移到一个新的盘子中,在盘子中可以汇集雏形以备后用(牵拉的玻璃杯移液器提供了一个简单的转移基础的方法:熔化玻璃的热量拉他们将消毒其末端)。
      注意:在整个解剖过程中,应注意确保介质不会长时间残留在空气中(介质中的pH指示剂会通过颜色变化显示出来),我们建议15分钟作为最大但是空气流动可能意味着需要更短的时间,根据我们的经验,HEPES抑制正常的肾脏发育,我们避免使用它)

  3. 塞宾格文化
    1. 将盖玻片和硅胶锥体放入80°C的烘箱中20分钟晾干(干燥对硅胶锥和盖玻片之间的适当粘合是必不可少的)。他们可以戴着手套的手或镊子使用塑料镊子来避免使用金属可能导致的损伤。
    2. 将一个锥形的窄端向下放在盖玻片上,用一个钝端的镊子将硅胶锥的边缘压在盖玻片上,以确保紧密粘附。
    3. 将带有硅胶锥的盖玻片放在60毫米培养皿的底部。 (图3)


      图3.赛宾格的培养方法A.显示培养体系的不同组成部分。 B.显示组装的赛宾格文化系统。

    4. 吸取一个肾脏雏形到由锥体定义的玻璃圆圈的大致中心。
    5. 为了避免组织变干的风险,应尽快吸除肾脏遗留的任何培养基,并将85μl培养基放入圆圈中。它可以直接从冷藏库中使用。使用移液器吸头确保这种介质扩散到覆盖整个由锥体包围的玻璃圆圈,并且不会“滴”成一个滴(图4)。请注意,最适量的介质会随着锥体的批次和玻璃的不同而略有变化。因此,我们建议用户用自己的材料尝试一系列的体积(,例如,80,82.5,85.97.5,90μl)以确定最佳效果。
    6. 将60毫米培养皿(盖玻片,硅胶圆锥体和肾脏雏形现在里面)放到100毫米培养皿的底部,并加入3毫升水合缓冲液(当水合缓冲液与硅胶锥体将锥体从盖玻片上分离,并且系统变得泄漏,这就是为什么我们使用单独的碟子来添加水合缓冲液)。 (图4)


      图4.圆锥和碗碟的排列:一种。在KCM培养基的少量培养基中,在硅胶圆锥体的中心培养肾脏原始成分,并在外部培养皿中加入增湿缓冲剂以防止干燥。 B. KCM倾向于作为一滴(左手侧图像),需要用枪头分布以覆盖由硅胶锥体(右侧图像)包围的玻璃环。

    7. 将盖子放在100mm培养皿上,然后将培养皿置于组织培养培养箱(37℃,5%CO 2,100%湿度)中。肾脏可以培养3天;除此之外,更改媒体是可取的。如图1A所示,外周的一些细胞可以出来并在玻璃上形成单层,但是整个肾脏应保持完整和连贯。如果“四舍五入”,但仍然活着,可能中间太多了;如果它变成一团干的,那就太少了。
      注意:对于延时拍摄,我们推荐Etaluma的孵化器内显微镜。
    8. 根据适用于任何抗体染色的方案,将标本固定在盖玻片上。除去培养基后,我们一般在-20°C下使用100%的甲醇,15分钟后使其升温至室温,然后用磷酸盐缓冲液洗涤。如果其他固定技术涉及洗涤剂(Tween,Triton等),应非常小心地更换溶液,以避免玻璃与培养物分离。在Bijou(5毫升)瓶中,抗体孵育通常在4℃过夜。用于图1中层粘连蛋白染色的实例方案是在4℃在PBS中的200μl1/100抗层粘连蛋白(Sigma-Aldrich)中过夜温育,在4℃下在PBS中7小时洗涤,过夜温育在4℃下在PBS中的200μl1/100 FITC抗兔(Sigma-Aldrich)中,在4℃在PBS中两次洗涤2次,然后在PBS或1: 1PBS-甘油,盖玻片与主玻片保持分离,盖玻片用作间隔物。这防止了组织的挤压。我们不嵌入或部分。

数据分析

  1. 这是一种文化的方法,而不是一个具体的测量方法,这种方法将支持一系列的问题和分析。但是,它的功能使其更适合于某些问题。为避免因个体差异造成的假阳性数据,如果要使用多于一只母鼠的胚胎,我们建议将所有胚胎的肾脏合并,然后随机分配给实验组和对照组。
  2. 该方法涉及的手工操作的程度使得器官雏形的最终尺寸和形状变化很大,并且一些根本不生长,这通常是因为如图4B左图所示,介质膜已经变成下降。至少,实验者需要解决一些排除标准,以便可以从分析中公平地排除失败的文化(理想的是,除了简单观察中等影片之外,在进行任何分析之前)。样本之间生长的变化,可能是由于样本在薄膜中的位置的细微差异而引起的,这意味着在实验和对照肾脏的任何有意义的比较之前,诸如总器官面积之类的特征的测量将产生大的误差条并且需要许多重复。我们建议不要使用这样的措施。更好的定量测量包括集合管树中分支尖端的数量和肾单位的数量。分支尖端计数分析的实例可以在Fisher等人(2001)和Michael等人(2005)中找到。即使如此,总是有一个模棱两可的问题:“T形”如何作为一个分支的终点必须被视为两个小窍门,而不是一个,而且对于什么是成熟的被称为肾单位,总是存在一些模棱两可的问题。因此,我们强烈建议所有的样本都是盲目编码的,或者如果这样做不实际,那么他们的图像是盲目编码的,并且由不知道样本是来自实验还是控制的人计数。
  3. 通过观察器官培养所获得的大量信息,无论是正常的还是受试验操作的,都是定性而不是定量的:管状形态是一个例子。同样,鼓励研究人员使用盲编码,也许使用由多个独立人员查看的图像,甚至可能使用半定量评分系统,甚至形态学(例如,将肾单位分类为“正常”或“异常”) 。一些数据是关于方向的,例如Henle环路的增长方向。方向需要坐标系统。可以通过从例如肾的中心(例如,使用ImageJ的质量函数中心确定的)径向地延伸线或者从第一分支点径向延伸线来制作简单的坐标系。例如Henle的圈的方向可以根据圈的细管与通过其末端的径向线之间的角度来测量。同样,在相同的盲编码样本上进行多个生物学重复和多个人测量将提供观察者间变异性和样本间变异性的良好测量。

食谱

  1. 清洁解决方案
    这由5:1:1(体积比)组成:H 2 O:H 2 O:2:NH 4 4 / sub > OH
    通过在水中加入过氧化氢和氢氧化铵,而不是相反,在处理浓缩溶液时使用个人防护装备(手套,眼睛保护装置)。
  2. 解剖介质
    根据生产商的说明,将Eagle's最小的基本培养基与厄尔氏盐和非必需氨基酸溶解在ddH 2 O中。
  3. 培养基:KCM
    注意:只需在Eagle's培养基中加入血清和抗生素(解剖介质)即可。
    解剖介质
    10%胎牛血清
    1/100青霉素/链霉素原液(该原液含有10.000U / ml青霉素和10mg / ml链霉素)
  4. 水化缓冲液
    PBS(来自Sigma-Aldrich片剂:让片剂溶解一小时,并充分混合,然后加入从上述方法3中提到的从100x储液中稀释的青霉素/链霉素)。

致谢

这里描述的技术的开发和使用得到了以下的资助:欧盟委员会FP6资助MRTN-CT-2006-036097,医学研究委员会资助MR / K010735 / 1,肾脏研究英国资助RP_002_20160223,以及埃及教育和文化局的博士奖学金。该协议最初在Sebinger,2010年的方法部分进行了描述。作者没有竞争的兴趣。

参考

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引用:Elhendawi, M. and Davies, J. A. (2018). Sebinger Culture: A System Optimized for Morphological Maturation and Imaging of Cultured Mouse Metanephric Primordia. Bio-protocol 8(4): e2730. DOI: 10.21769/BioProtoc.2730.
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