小鼠ES细胞核型分析

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http://www.millipore.com/userguides/tech1/mcproto0374 [  z( q& z0 E/ U
Protocol: Karyotyping ES Cells8 R, e2 j9 f- x, r- ^5 ]( E
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Catalogue Number:        mcproto0370 N0 y6 q1 @& |* q' p0 P" e0 |. `

9 K' t; [* D+ q# KYear:        2007       
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; N, G7 I5 E2 V' r* sThis method works best with actively growing culture of ES cells (i.e. 1-2 day culture).
* U2 K% B5 C0 t# X6 L! g( @Method
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1.        The day before, passage a 70% confluent ES cell plate 1:2.
1 |: z$ j. I- M2.        On the morning of karyotyping, change the medium on the plate at least 3 hours before passage and collection.
1 D5 E2 K1 q. \( P3.        Trypsinize and collect ES cells into a conical tube. Centrifuge as usual and aspirate medium. Avoid allowing the pellet to dry out.
( X  |; r- J; w' m& Q0 r4.        Gently flick the tube to resuspend the cell pellet, and add 8mL of hypotonic KCl solution to the cells. Continue to gently flick the tube during the addition of KCl to avoid clumping.; U# d2 R; z$ C; C: y% S8 O
5.        Incubate the tube at 37°C for 10 minutes (this may vary for each type of cell line used).
* z% ~* b, b: [' H! q6.        Add 2mL of freshly made fixative and mix by gentle inversion. {Fixative MeOH:Glacial Acetic acid 3:1 made fresh and stored at 4°C}.
! u6 ?  \4 Z' Z! {: m% Q7.        Centrifuge cells at 1000 rpm for 5 minutes and aspirate supernatant.
9 E+ v2 h# O% t) B8 U8.        Using a pasteur pipette, carefully add 2mL of fixative solution dropwise, with gentle mixing to avoid clumping. Add an additional 6mL of fixative and mix by gentle inversion of the tube.
% }$ B: Y, t4 r3 G* y: {. Y% u9.        Centrifuge cells at 1000 rpm for 5 minutes and aspirate supernatant.  o$ L2 W% l1 j# s
10.        Repeat steps 8 & 9 three times.
. M, L, h! X" N, k8 S" S; g11.        Resuspend the pellet in 1mL of fixative (less or more according to pellet size).7 D8 p$ c+ H# W8 e3 K

( v, ~# K6 i+ _* L4 f, O2 R" OTo make cell spreads, firstly humidify the surface of a dried cold slide by application of warm breath, whilst holding the slide at a 45° angle. Using a pasteur pipette, carefully drop (from a height of approx 0.5 metres) one drop of the suspended cells onto the top surface of the slide and allow to air dry.
1 A5 B% I. Y6 `4 x! EStaining3 u$ S% @/ Q% b- l

. ]) o. i) Q+ N/ }9 nStain slides with freshly made Leishmann’s stain for 8 minutes.
" Y# w& g* @# {( c' D: \. C( Q: ARinse in running water for 1 minute and air dry.
  N5 M8 c9 G) F2 oClear cells in 2x changes of xylene and mount coverslip using Depex.
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: e9 ^2 l4 q1 C8 U  p6 gColcemid is not used in this method, as the mitotic index of actively growing ES cells is generally high enough to get a good chromosome spread.# k! G0 j# a0 {; V( q
High quality slides need to be used. Slides should be soaked in 100% ethanol overnight and dried with lint-free tissue before use. As it is important to have slides “cold” before use, slides in ethanol bath can be stored in fridge or freezer until ready to make cell spreads.5 x; @. l6 r  P- S* w6 ~
Some notes on KCl:- Most labs use 0.56 % KCl and some labs use 0.2% KCl + 0.2% Na citrate instead. This depends entirely on the cell types being analyzed. The time in KCl is crucial – too short and the chromosomes will be too tightly packed; too long and they will not remain in their appropriate group.

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G Banding- m$ n) `: L  g
10. Place suitably aged slides (see note below) in 2X SSC in a Coplin staining jar with a lid in a water bath at 60°C-65°C for 1.5 h. Then cool the slides to room temperature by running tap water over the closed jar. Transfer the slides to 0.85% (w/v) NaCl at room temperature for 5 min.
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Before G banding, slides should be “aged” for between 3 and 21 d by leaving them in a closed box at room temperature. Fresh slides give poor G-band resolution. Maximum G-band resolution is achieved at ~10 d after slide preparation. Beyond this time, resolution slowly decreases until, after several weeks in storage, the chromosomes either fail to band and stain uniformly or show “pseudobands” that are not significant to the standard idiogram.
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11. Drain the slides by touching them onto filter paper. Place them on a flat surface and flood the chamber with 0.025% trypsin in 0.85% NaCl for 15-20 sec.
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3 ?- ~$ E3 k4 u7 k8 gThe trypsin exposure time is critical: Underexposure preserves chromosome morphology but gives poorly differentiated bands, and overexposure distorts morphology and eliminates most of the bands. Optimum trypsin times are known to vary among laboratories. A test slide should be treated for the minimal suggested time of 15 sec to establish the best treatment time for the rest of the slides.: G- ]( o9 K# N% O9 W5 d9 o/ d

& q2 Z* q4 h" p4 g0 r5 ~In the laboratory of E.P. Evans, the optimal trypsin exposure time has been established as between 15 and 20 sec for mouse chromosomes, irrespective of the source of the mitotic cells./ T" d5 N3 M7 |1 d" ]2 v
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12. Stop tryptic activity by placing the slides back into 0.85% NaCl. Then rinse slides in phosphate buffer (pH 6.8) and stain in fresh Giemsa stain in 5 mM phosphate buffer (pH 6.8). After 10 min in the stain, monitor wet slides under low-power, bright-field microscopy (160X) for staining intensity. Because, upon drying, wet slides gain contrast, care should be taken not to overstain the cells as this will reduce G-band differentiation. If necessary, repeat staining until adequate results are achieved and then quickly rinse slides in phosphate buffer (pH 6.8) and blow-dry with a current of cool air.' k- D# A/ C+ U9 g$ W. h2 g2 M( M
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13. Examine unmounted slides with a 100X oil-immersion lens.% O+ ~6 O  @- `3 @1 m
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The majority of modern, readily available immersion oils that are declared “PCB-free” also have the unfortunate property of removing Giemsa stain after a few hours of exposure. Although direct viewing of slides under an oil immersion lens gives a higher optical resolution, it is wise to mount slides if they are to be kept.