Materials and Reagents

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 )
40 x 0.13 mm borosilicate glass coverslips (VWR, catalog number: 631-0177 )
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.
100 mm sterile Petri dishes (for dissection–surface quality is irrelevant) (Cell Star^®, Greiner Bio One International, catalog number: 664160 )
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.
Glass Pasteur pipettes (150 mm, Volac, catalog number: D810 )
Timed-mated mice at E11.5 (E10.5 is also suitable but is a more challenging dissection)
Sterile distilled water
100% methanol
Eagle’s minimal essential medium with Earle’s salts and non-essential amino acids (Sigma-Aldrich, catalog number: M5650 )
Foetal bovine serum (FBS) (Biochrom, catalog number: S 0415 )
Penicillin-streptomycin (Sigma-Aldrich, catalog number: P4333 )
Phosphate buffered saline (PBS) (Sigma-Aldrich, catalog number: 79382 )
Anti-laminin (Sigma-Aldrich, catalog number: L9393 )
FITC anti-rabbit (Sigma-Aldrich, catalog number: F0382 )
EmbryoMax^®Penicillin-streptomycin solution, 100x (Sigma-Aldrich, catalog number: TMS-AB2-C )
Hydrogen peroxide (Sigma-Aldrich, catalog number: H1009 )
Ammonium hydroxide (Sigma-Aldrich, catalog number: A6899 )
Cleaning solution (see Recipes)
Dissecting medium (see Recipes)
Culture medium (see Recipes)
Hydration buffer (see Recipes)

Equipment

Forceps
Scalpel (curved blade, e.g., D-form, type 22, Swann Morton, catalog number: 0108 )
80 °C oven
Note: We use a Gallenkamp Hotbox size 1, but any 80 °C oven should be suitable.
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.
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.
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

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.
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).
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

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.
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.
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

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
Dissecting medium
Dissolve Eagle’s minimal essential medium with Earle’s salts and non-essential amino acids in ddH₂O as per manufacturer’s instruction
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)
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

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