Stripping Western Blots
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1) After ECL development, wash membrane once for 10min with PBST.
2) Incubate the membrane in stripping buffer (see below) in a heat-sealed plastic bag for 30min at 50oC with occasional mixing. For low-abundance antigens, the strip can be done overnight at RT with constant agitation.
3) Washed stripped membrane at least 3x for 10min each with PBST, then re-block (in PBST+NFDM or PBST+BSA) overnight at 4oC.
Stripping Buffer: 62.5mM Tris pH 6.8
2% SDS
100mM b-ME
Quick Mix: Use 4X stacking buffer at 1:8, 10% SDS at 1:5, and b-ME at 70μl/10ml
(e.g.10mls strip=1.25mls 4X, 2mls 10% SDS, 70μl b-ME & 6.68mls H2O)
Lysis of Cultures Cells General Protocol
1) Wash cells twice with ice-cold PBS. Use 3mls (6cm plate) to 5mls (10cm plate) per wash. (For cells in suspension, pellet cells at room temperature, aspirate & discard mediate, add 5-10mls PBS, resuspend with 1ml pipetman, and spin for 2-3min at 1500rpm in refrigerated centrifuge.)
2) Aspirate PBS, let plates drain on an incline of ice for 15", then aspirate off remaining PBS.
3) Add appropriate lysis buffer. Use 0.5-1ml per 10cm plate and 0.25-0.5ml per 6cm plate.
4) Tilt plates back & forth a few times (to ensure even spread of buffer) and place on ice or in refrigerator for 10min.
5) Scrape into pre-chilled microfuge tubes, vortex for 10-20", and place on ice for 10min. (For cells in suspension, add lysis buffer to pelleted cells, triturate with pipetman, and ice for 10min, then transfer to microfuge tube, vortex, and ice for another 10min.)
6) Spin at 14000rpm (~16000xg) for 10min at 4OC.
7) Transfer supernatant to new, pre-chilled microfuge tube.
8) Use 5-10 μl for protein assay, or mix 20 μl with 20 μl 2X sample buffer (or 30 μl with 10 μl 4X sample buffer) for western blotting. Store the rest at -80OC.
Lysis Buffers and their Applications
RIPA Buffer - For high-stringency lysis (disrupts most protein-protein interactions)
1% NP40
0.5% NaDOC (deoxycholate)
0.1% SDS
50mM Tris pH 7.4-7.5
150mM NaCl
10% glycerol
1mM EDTA
Modified RIPA Buffer - Good general-purpose buffer
1% NP40
0.5% NaDOC
50mM Tris pH 7.5
150mM NaCl
5mM EDTA
NP40 Buffer - Better than mRIPA for some protein-protein interactions but not as clean
1% NP40
50mM Tris pH 7.5 (Note: can increase to pH 8 to decrease background)
150mM NaCl
5mM EDTA
Triton X100 Buffer #1 - Use for PAK-Nck and some other co-IPs
1% Triton X100
50mM Tris pH 7.2
10% glycerol
25mM b-glycerophosphate
2mM EDTA
2mM EGTA
Triton X100 Buffer #2 - Use for high-stringency preparation of cytoskeleton (pellet)
1% Triton X100
0.27M sucrose
20mM Tris pH 7.2
10mM b-glycerophosphate
1mM EDTA
1mM EGTA
0.1% b-mercaptoethanol
For most applications, use the following protease and phosphatase inhibitors ([final;stock]): NaF (1mM; 1M), Na3VO4 (1mM; 0.1M), AEBSF (100μM; 100mM), benzamidine (5mM; 1M), aprotinin (0.1% of Sigma A6279).
Can also add nitrophenyl phosphate (1mM; 0.1M) and calyculin A (20nM; 100μM) to further inhibit phosphatase activity, if necessary.
Immunoprecipitation General Protocol
1) Wash cells twice with PBS and lyse in appropriate lysis buffer;
Dish size Wash vol. Lysis vol.
1 well 1ml 125-250μl
6cm plate 3mls 250-500μl
10cm plate 5mls 0.5-1ml
2) Incubate on ice for 10min. Scrape into microfuge tube, vortex for 10”, then ice for an additional 10min.
3) Spin at 4OC for 10min at 14000rpm (Eppendorf microfuge; ~16000xg)
4) Transfer supernatant lysate to new, pre-chilled microfuge tube and remove 5-10μl lysate for protein assay. Discard pellet or, if desired, resuspend in 50-200μl of 1X sample buffer, sonicate and boil.
5) If desired/necessary, pre-clear the lysate by adding 30μl protein A-, protein-G, or protein A/G-sepharose (at 1:1 slurry) and incubating for 30min at 4OC with rocking.
6) Spin at 4OC for 2min at 14000rpm.
7) Transfer lysate, leaving the last 10μl over the pellet (if possible), to another new, pre-chilled microfuge tube containing an appropriate amount of desired antibody. The amount will depend on the antibody and the abundance (total and relative) of the antigen, but a good starting point is 0.5μg antibody / 500μg lysate.
8) Incubate for 1-2hr at 4OC with rocking. Avoid overnight incubationsa.
9) Add 30-40μl protein A-, protein G-, or protein A/G-sepharose and incubate for 30min to 1hr at 4OC with rocking.
10) Collect beads by spinning at either 14000rpm for 10-15” or 5000rpm for 1min (if the latter, spin at 4OC). Carefully aspirate supernatant from beads. Add 1ml lysis bufferb and vortex briefly.
11) Repeat Step #10 2-3 more times.
12) After last wash, aspirate supernatant and remove remainder of wash buffer with fine gauge (>22g) needle. Process for kinase reaction (as indicated) or add 40-80μl 1X sample buffer and boil for 5min.
Notes: a If antigen is in low-abundance, premix antibody and protein (A/G)-beads, in lysis buffer, for 1hr prior to addition to pre-cleared lysates. This increases the avidity of the immunocomplex but does not increase background.
b Some immunoprecipitations will benefit significantly from following the first or second lysis buffer wash with 1-2 high salt washes (e.g. 500mM LiCl/100mM Tris pH8.6) followed by a low-salt wash (e.g. a 1:5 dilution of high-salt wash). Also, if immunoprecipitating for a kinase reaction, the last wash or two should be in the appropriate kinase buffer.
Immunoprecipitation Kinase Assay MAPK
1.) Wash cells 2x with ice-cold PBS.
2.) Add RIPA lysis buffer (0.1% SDS, 0.5%DOC, 1.0% NP40, 50mM Tris pH 7.4, 150mM NaCl, 1mM EDTA) containing phosphatase and protease inhibitors (I use PMSF, aprotinin, pepstatin, NaF and Na-orthovanadate). Use 0.5-1.0ml lysis buffer for 10cm plate (depending on cell density), and scale up or down accordingly.
3.) Scrape lysate and transfer to microfuge tube.
4.) Vortex (10sec) and incubate on ice for 10min.
5.) Spin in a microfuge for 10min at 4oC at 14krpm.
6.) Transfer supernatant lysate to new microfuge tube. Freeze at -80oC or proceed directly to IP. At this point you can pre-clear the lysate by incubating with 30μl protein A/G sepharose for 30min at 4oC with rocking, then centrfuging in the cold for 1min at 14krpm and transfering the supernatant lysate to a fresh tube. Depending on the cell type you are using, this may or may not be necessary - however, it never hurts to do it.
7.) Determine protein concentration by whatever method you’re used to.
8.) Typically, I use 150-200μg per immunoprecipitation. You can use less (as little as 50) but the higher amount gives you a good, easily detectable amount of activity.
9.) To the lysate, add 1.5 μg of anti-ERK antibody (Santa Cruz, SC154 (Erk2) or SC93 (Erk1)), and incubate, with rocking, at 4oC for 1hr.
10.) Add 20-30 μl of protein A/G sepharose (50% slurry) and incubate at 4oC for 30min to 1hr.
11.) Prepare kinase buffer (50mM HEPES or Tris pH 7.4, 10mM MgCl2, 10mM MnCl2, 1mM DTT) and reaction mixture, which is 15 μM ATP, 500 μg/ml MBP in kinase buffer.
12.) Wash the immunoprecipitate 3x with RIPA buffer and once with kinase buffer.
13.) Add 40 μl of the reaction mixture, along with 15 μCi of gamma-32P-ATP (3000Ci/mmol, NEN) to the washed beads and mix very gently by tapping the tube (try to avoid getting beads stuck to the side of the tube, above the level of the reaction mixture).
14.) Incubate at 25oC (room temp.) for 25min, mixing occasionally.
15.) Add sample buffer to 1x (I usually add 20 μl of 3x), and boil samples for 3-5min.
16.) Run samples on 12% or 15% gel. Stain gel, dry and expose to film or Phosphorimager screen. I usually load between 20 and 30 μl, and I can see a strong signal overnight. Sample can also be stored, after boiling, at -20oC.
IPKA HA-tagged MAPK (a variation & alternate protocol for MAPK IPKA)
1) Transfect cells with GFP and pcDNAI-ERK1-HA plasmids
2) Transfected cells were allowed to adhere to fibronectin-coated dishes for the given time and then stimulated accordingly with growth factor.
3) Wash cells x 2 with ice-cold PBS and then lyse in modified RIPA buffer (50 mM Hepes, pH 7.5, 1% NP-40, 0.5% sodium deoxycholate, 150 mM NaCl, 50 mM NaF, 1 mM sodium vanadate, 1 mM nitrophenylphosphate, 5 mM benzamidine, 0.2 μM calyculin A, 2 mM PMSF, and 10 μg/ml aprotinin).
4) Lyse for 20 min on ice and clarify lysates by centrifugation at 16,000 x g for 10 min at 4 oC
5) Preclear lysates with 30 μl of a 1:1 slurry of protein G-sepharose (Fast-Flow; Pharmacia Biotech, Cat# 17-0618-01) on a rotator at 4 °C for 30 min.
6) Incubate precleared lysates with anti-HA antibody on ice for 2 h.
7) Add 30 μl Protein G beads and incubate on a rotator at 4 °C for 2 h.
8) Wash immunocomplex once with cold lysis buffer.
9) Wash twice with cold wash buffer (0.1 M NaCl, 0.25 M Tris-HCl, pH 7.5.).
10) To washed immunocomplex, add 40 μl of reaction mixture (10 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 1 mM DTT, 25 μM ATP, 5 μCi 32P-ATP (Dupont NEN Cat# NEG-002A) and 10 μg myelin basic protein (UBI or GIBCO)) per assay and incubate on a shaking platform at RT for 30 min.
11) Add 13 μl 4 X SDS sample buffer and boil for 5 min to stop the reaction.
12) Separate reaction by SDS PAGE: 15% to analyze incorporation of 32P into myelin basic protein and 10% gel for Western blotting to obtain levels of expressed HA-ERK1.
IPKA MEK
1) IP MEK from 200-400μg of lysate using 1-2μg of the Transduction Laboratories anti-MEK1&2 antibody (catalog number M17030)
2) Wash IP 3X with lysis buffer and once with 1X kinase buffer (50mM HEPES or Tris pH 7.4, 10mM MgCl2, 10mM MnCl2, 1mM DTT)
3) Prepare Reaction Mix (use 40μl per IP):
(per IP) 20μl 2X Kinase Buffer
0.6μl 1mM ATP (final [ ] = 15μM)
5-10μl k- MAPK
0.5μl 32P-ATP
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4) Incubate at RT for 20-25min or at 30oC for 10-15min.
5) Stop reaction with appropriate amount of concentrated sample buffer (e.g. 40μl of 2X).
6) Run 20-40μl on 10 or 12% SDS-PAGE gel, stain, dry and expose.
IPKA - PAK
1. Wash cells twice with ice-cold PBS, drain well, then add any of the following buffers (containing protease & phosphatase inhibitors): modified RIPA, NP40 lysis buffer, Triton X100 lysis buffer. Use ~750μl/10cm plate (500μl/6cm, 250μl/well of 6-well plate).
2. Spread lysis buffer evenly over cells, let plate sit on ice for 5-10min, then scrape lysate into pre-chilled microfuge tube.
3. Vortex lysate vigorously for 10sec, incubate on ice for 10min, then spin at 14000rpm for 15min at 4oC.
4. Transfer supernatant to new, pre-chilled microfuge tube and determine protein concentration.
5. Incubate equal amounts of lysate (> 500 μg; the more the better) with 0.25-0.5 μg anti-PAK antibody per 100 μg of lysate for 1.5hrs at 4oC with gentle rocking.
6. Add 25 μl protein A-, protein G-, or protein A/G-beads and incubate for 30min at 4oC with gentle rocking.
7. During this incubation, make the kinase buffer. Final concentrations for 1X are:
25mM Tris (or HEPES) pH 7.5
10mM MgCl-2
5mM MnCl2
0.2mM DTT
To make the kinase buffer, add all of the components and bring to 1/2 the final volume (e.g. if making 10mls total, mix the components and bring to 5mls with H2O), then mix well and remove and reserve 0.5mls (this is 2X kinase buffer). Add an equal volume of water (e.g. 4.5mls) to the remaining buffer to bring it to 1X concentration.
8. Wash beads 2 times with lysis buffer (including phosphatase inhibitors in the washes can help preserve maximum activity), once with lysis buffer containing 500mM NaCl, and once with 1X kinase buffer. Carefully remove as much of the final wash buffer as possible.
9. The reactions are carried out in 40 μl 1X kinase buffer containing 0.1mg/ml (1 μg/10 μl or 4 μg/reaction) myelin basic protein, 50 μM ATP, and 5-10 μCi 32P-ATP. To set this up, for each reaction, combine 20 μl 2X kinase buffer, the additional reaction components (MBP and ATPs), then bring to 40 μl total (per reaction) with water. It is best to make enough for one or two extra reactions to ensure having enough volume for each sample, plus a reaction-only control.
10. Incubate reactions at 30oC for 30min, then add an equal volume of 2X sample buffer (or 10 μl of 5X or whatever you prefer) and boil the reactions for 5min. Run > half of the reaction on a 12% or 15% gel, stain with Coomassie, destain, dry and expose to film or phosphorimager screen.
Abl Kinase Assay Protocol (from R. Plattner & A.M. Pendergast)
1. Following stimulation, place cells on ice and wash 2X with cold PBS
2. Lyse each 100 mm plate with 500-1000 μl of lysis buffer
(50mM HEPES pH7.0, 150 mM NaCl, 10% glycerol, 1% triton-X-100, 1.5
mM MgCl2, 1mM EGTA and protease and phosphatase inhibitors)
Protease inhibitors―add fresh Aprotinin 10 μg/ml
Leupeptin 10 μg /ml
Pepstatin 10 μg /ml
PMSF 1 mM
NaF 25 mM
Na Orthovanadate 1mM-make that day
3. Scrape cells off of the plate, put in an eppendorf, rock 10-20’ at 4C and then
spin 10’. Save supernatent―determine protein concentration
4. Immunoprecipitate 100 μg of protein (if you use K12 antibody, and 200 μg of
protein if you use AB-3 antibody) in a total of 800 μl. Use 1 μg of antibody
per reaction.
5. Add protein A sepharose or protein G sepharose depending on which
antibody used to IP.
6. Wash complexes:
1: 2X with 0.5 ml of RIPA buffer (50 MM Tris pH 7.5, 150 mM NaCl, 1%
triton-X 100, 0.1% SDS, 1% sodium deoxycholate) + inhibitors
2. 2X with 0.5 ml of buffer containing 10 mM Tris, pH7.4, 5 mM EDTA, 1%
triton-X-100, 100 mM NaCl + inhibitors
3. 2X with 0.5 ml of buffer #2 without NaCl
4. 2X with 1 ml in kinase buffer containing 20 mM Tris pH7.4, 10 mM
MgCl2, 1 mM DTT
The kinase reactions are performed in a volume of 20 μg containing 1 μM cold ATP and 5 μCi gamma-32P-ATP and 0.5 μg GST-Crk in kinase buffer. Incubate the kinase reactions for 40 min. at room temp. Stop reaction by adding 20 μl of 2X SDS-sample buffer. Always check that the assay is in the linear range otherwise differences will be muted, particularly if using overexpressed c-Abl. The previous protocol is for endogenous c-Abl. Finally, be aware that K12 immunoprecipitates a serine kinase in addition to c-Abl in some cell lines such as MEFs and so can’t be used reliably in those cells.
In-gel Kinase Assay Protocol #1 7-10-2001
1. Pour appropriate percentage gel containing ~50μg/ml of peptide substrate or ~0.5mg/ml protein substrate.
2. After electrophoresis, wash gel in SDS Removal Buffer for 1hr with three changes of buffer, then with Buffer A for 1hr.
3. Incubate gel in Denaturation Buffer for 1hr.
4. Rinse gel 2-3 times with Renaturation buffer, then incubate in Renaturation buffer overnight at 4oC.
5. Bring gel up to room temperature in Renaturation Buffer, then rinse and wash for 1hr with Kinase Buffer.
6. Incubate gel in Kinase Buffer containing 2-5μCi/ml of 32P-gamma-ATP for 3hrs at room temperature (> 20oC).
7. Wash gel extensively (1-2hrs with > 4 changes of buffer) with Stop Buffer.
8. Dry and expose to film or phosphorimager plate.
・ 2X Buffer A: 100mM Tris (pH 8.0), 10mM 2-ME
・ SDS Removal Buffer: 20% isopropanol in 1X Buffer A
・ Denaturation Buffer: 6M guanidine-HCl in 1X Buffer A
・ Renaturation Buffer: 0.04% Tween 20 in 1X Buffer A
・ Kinase Buffer: 25mM Tris (pH 8.0), 2.5mM 2-ME, 10mM MgCl2 (10mM MgCl2 in 0.5X Buffer A)
・ Stop Buffer: 1% sodium pyrophosphate in 5% TCA
In-gel kinase assay Protocol #2[1]
1) Add substrate to separating gel to a final concentration of 0.1mg/ml (for peptides) to 0.5mg/ml (for MBP or fusion proteins) and polymerize as usual.
2) For whole cell extracts, run 10-50μg protein per lane, or run 50-100% of an IP.
3) Wash gel twice for 30min each with 150-200mls 20% isopropanol in buffer A (50mM HEPES pH 7.4, 5mM b-ME)[2].
4) For denaturation, incubate the gel for 1h in ~150mls 6M guanidine HCl in buffer A. If denaturation is not needed/desired, skip to step 5.
5) For renaturation, wash the gel extensively[3] with 0.04% Tween 20 in buffer A at 4oC.
6) Wash gel once for 30min at 30oC in kinase buffer (buffer A supplemented with 10mM MgCl2).
7) Incubate gel for 30-60min at 30oC in kinase buffer containing 50μM ATP and 5-20μCi/ml of gamma-32P-ATP.
8) Remove kinase mixture and wash the gel several (5-8) times for 15min each at room temperature with 5% (v/v) trichloroacetic acid and 1% (w/v) sodium pyrophosphate, removing the first two washes to liquid radioactive waste. If background is still high, wash gel once more overnight at 4oC.
9) Dry and expose to film or phosphorimager plate.
Guanidine HCl = 95.53g/mol = 47.76g/100ml for 6M
Sodium pyrophosphate = 446.1g/mol (1% = ~ 20mM)
b-mercaptoethanol = 14.3M stock = 349μl / l for 5mM
TCA (trichloroacetic acid) precipitation of proteins
1) It is somewhat difficult to work with volumes of less than 100μl. Therefore, if your sample is more concentrated than that, dilute it to > with water or, better still, the buffer it is already in. Also, you can precipitate as little as 10μg and get a workable pellet, but it is small. Precipitation of at least 25μg is recommended.
2) On ice, pre-chill sample, microfuge tubes, and enough acetone to do 2x 0.4ml washes per sample.
3) Add sample to chilled tubes and dilute as needed.
4) Add 100% (w/v) TCA to the samples to a final concentration of 10%. For example, for a 100μl sample, the formula is 100μl + 0.1X = X (where 0.1 represents the 10% final TCA conc), or X=11.1 μl.
5) Mix well by inverting 5-6x, then place on ice for > 30min. This step can go overnight, as long as the samples are kept ice-cold.
6) Spin the samples in a microfuge in the cold at 16,000xg for 15min. There should be a fairly compact white pellet at the bottom of the tube.
7) Carefully aspirate the supernatant, using a loading tip on the end of a vacuum-linked Pasteur pipette. It’s better to leave some residual TCA than to aspirate your pellet.
8) Add 0.4ml ice-cold acetone to the samples (it is not necessary to vortex) and spin again in the cold at 16,000xg for 10min.
9) Carefully aspirate the supernatant.
10) Repeat Steps 8 & 9 once more, then let the pellet air-dry for 10-15min at RT. Take care to prevent dust & detritus from falling into the tube, especially if the sample is ultimately to be used for mass spectrometry.
11) Resuspend pellet in a buffer compatible with the intended next step (e.g. rehydration buffer for IEF, 0.1M Tris base + 0.1% SDS for protein assay, etc).
Coomassie Staining of Gels for Mass Spectrometry
1) Fix gel for 20min in 10% acetic acid/ 25% isopropanol. Wash once with deionized water.
2) Stain gel in 0.006% Coomassie Brilliant Blue R-250 in 10% acetic acid for 1-6 hrs. (0.006% = 30mg in 500mls)
3) Destain in 10% acetic acid 2-12hrs. If necessary, gel can be stored in 10% acetic acid at 4OC for up to 1wk until ready for in-gel digestion.
3 Bacterial Culture
Transformation of Competent E. Coli Cells 09/05/96
1) Thaw competent cells on ice. Throughout the procedure, do not allow cells to warm above 4oC and never vortex them. (The procedure that makes bacterial cells competent, by design, porates them and makes them fragile).
2) To a pre-chilled (and sterile, if possible) microfuge tube, add 0.1-10ng of DNA. The amount will depend on its origin (use ~0.1ng if transforming purified supercoiled DNA, and ~10 or more ng for ligated plasmids).
3) Add 50-100μl of competent cells, depending on the supplier, cell type, and DNA origin. (Use less for supercoiled plasmids, more for ligated plasmids). Mixing is not necessary and may lower efficiency.
4) Incubate on ice for 45-60min. During this time, pre-heat a water bath to 42oC.
5) Heat-shock the bacteria at 42oC for 1min, then replace the tube on ice for 1-2min.
6) Add 900-950μl LB or SOC medium to the cells and incubate at 37oC for 1h with gentle shaking. During this time, place in the dry 37oC incubator the appropriate number of LB-agar plates containing the appropriate concentration of the appropriate antibiotic(s) and allow to warm for 15-20 min.
7) Plate 5-100% of the culture onto an LB-agar plate. If plating > than 0.1ml, gently pellet the bacteria (e.g. 5 min at 500xg) and gently resuspend (by pipetting, not vortexing) in ~0.1ml LB, then plate.
8) Let the plates dry at RT for ~5-10min, then invert and place at 37oC in a dry incubator.
9) Colonies should be present within 12h, but incubation can go for up to 24h. Alternatively, the plate can be kept on a clean bench top for ~48hrs (e.g. over a weekend).
Note: Snap-cap Falcon tubes (e.g. 2059’s) can be used instead of microfuge tubes. If so, heat shock for only 45”.
4 Cell Culture
Suspension Culture
General Considerations: The removal of the cells from their substrate should be as rapid as possible, to minimize exposure to trypsin. If a loosely adherent cell line (e.g. HEK293) is used, trypsin can be replaced by versene or EDTA alone, and the soybean trypsin inhibitor can be eliminated from the subsequent step. The bovine serum albumin (BSA) used should be delipidated and globulin-free to minimize the introduction of stimulatory substances with the suspension medium. As with all cell culture, avoid bubbles and harsh vortexing.
1. Serum-starve cells as appropriate (e.g. O/N for REF52 or NIH3T3,<12h for WI38)
2. Remove medium, rinse once with 0.05% trypsin/EDTA (e.g. 4mls/10cm plate), add fresh trypsin/ EDTA (e.g. 1ml/10cm plate), and incubate at 37oC as appropriate to loosen cells (e.g. 1-2min for NIH3T3, 2-5min for MDCK).
3. Add 4 volumes of soybean trypsin inhibitor (1mg/ml in serum-free medium) e.g. 4mls to 1ml of trypsin/EDTA and triturate to completely remove cells.
4. Collect cells by centrifugation in a conical tube at 500xg for 3-5min (depending on volume) at RT or 37oC.
5. Completely remove supernatant and gently resuspend cell pellet in media + 1% tissue culture grade BSA (e.g. A9306 from Sigma). The resuspension volume will depend on the intended usage and desired plating density of the cells, and should average 1x106 cells per ml or less. ***If desired, another round of centrifugation, decanting, and resuspension can be performed to more thoroughly wash the cells, but for most applications this appears unnecessary.
6. Place tube on Nutator (or the like) in a 37oC incubator for the desired time.
PC12 Cell Culture
Coating plates
1) Poly-D-Lysine (Sigma # P7280)
Prepare frozen stock in ddH2O @ 100 μg/ml
Freshly dilute to 25 μg/ml and coat at 5 μg/cm2 (5 ml/25 cm2)
Coat overnight at 37 °C.
Wash with sterile PBS
2) Collagen, Type I Rat Tail (Sigma # C7661)
Prepare 4 °C stock in 0.25% acetic acid @ 1 mg/ml
(takes several hours to dissolve, remove any undissolved material by centrifugation)
Dilute to 50 μg/ml in PBS & coat at 10 μg/cm2 (5 ml/25 cm2)
Coat 2 h at 37 °C
Wash with non-serum containing medium
Culture Media
85% RPMI 1640
10% Heat-inactivated horse serum
5% Fetal bovine serum
Glutamine
Differentiating Medium
92.5% RPMI 1640
5% Heat-inactivated horse serum
2.5% Fetal bovine serum
2 nM Nerve growth factor (Boehringer Mannheim #1058 231)
Subculture Split at 1:3 to 1:4 once a week
Partial media change every 3 days
Differentiation Plate cells so cells are dispersed
After 7 days NGF treatment, most cells have processes
Maximal after 14 days
Transfection of Fibroblasts - SuperFect
1) Seed cells in mid- to late afternoon and let grow overnight.
NIH3T3 2x105/well of 6-well dish
6x105/6cm plate
REF52 1x105/well of 6-well dish
2.5x105/6cm plate
WI38 3x105/well of 6-well dish
2) In a sterile microfuge tube, prepare the following:
Per well of 6-well dish 90μl OptiMEM
2μg DNA (total)
8μl SuperFECT reagent
Per 6cm plate 130μl OptiMEM
6μg DNA (total)
20μl SuperFECT
Vortex briefly & gently (~50% power) and incubate at room temperature for 20min.
3) Add complete media (DMEM+10% CS (NIH3T3, REF52) or MEM+10%FBS (WI38)) to each tube.
0.6ml / tube for 1 well
1.0ml / tube for 6cm plate
4) Remove media from cells. Triturate transfection mixture to mix, then gently pipet onto cells.
5) Incubate 4hrs at 37OC.
6) Remove transfection solution from cells and replace with fresh, complete media.
7) Depending on experiment, harvest 24-48 hrs post transfection.
LipofectAMINE Transfection of NIH3T3 Cells
1. Plate 3T3 cells at 2 x 105 per 35 mm dish & let grow overnight (cells should be 50-70% confluent for transfection).
2. For each transfection, dilute:
A: 8 μl LipofectAMINE in 100 μl OptiMEM
B: 2 μg DNA in 100 μl OptiMEM
3. Mix A and B, incubate for 30 min at room temp for DNA-liposome complexes to form.
4. Wash cell monolayers once with OptiMEM, then add 800 μl OptiMEM to cells in each 35 mm well and then add Lipofectamine/DNA mix.
5. Incubate for 6 h, then add 4 ml of DMEM/FBS per well.
6. Replace with fresh media after 24 h, harvest cells 48-72 h after start of transfection.
LipofectAMINE - GIBCO BRL Cat# 18324-012
Preparation of Vitrogen Collagen Gels
(Protocol from Cohesiontech.com (supplier of Vitrogen ColI solution; Cat# FXP-019))
A. Materials
1. Vitrogen Collagen, chilled to a temperature of 4°-6°C.
2. Sterile 10X phosphate-buffered saline solution (0.2 M Na2HPO4, 1.3 M NaCl, pH = 7.4).
3. 0.1M HCl.
4. 0.1M NaOH.
5. Phenol red or pH paper.
B. Preparation of neutralized, isotonic Vitrogen Collagen solutions
1. Mix 8 ml of chilled Vitrogen Collagen with 1 ml of 10X phosphate-buffered saline solution. (Alternatively one can use a 10X solution of buffered cell culture media.) Add 1 ml of 0.1 M NaOH and mix.
2. Adjust the pH of the solution to 7.4 ± 0.2 by the addition of a few drops of 0.1M HCl or 0.1 M NaOH. The pH of the solution can be monitored by pH paper or by the use of a pH indicator (dye) such as phenol red. Phenol red can be added to the phosphate-buffered saline solution at a concentration of 0.005 mg/ml.
3. The neutralized, isotonic Vitrogen Collagen solution can be stored at 4°-6°C for several hours prior to gelation.
C. Gelation of neutralized, isotonic Vitrogen Collagen solutions.
1. Collagen gelation (fibrillogenesis) can be initiated by warming the neutralized Vitrogen Collagen solution to 37°C. Since gelation occurs more rapidly in the absence of CO2, we suggest that CO2 incubators not be used. For best results, allow a minimum of 60 minutes for gelation to occur.
2. Cells can be dispersed on collagen gels, sandwiched between collagen gels or suspended in collagen gels by mixing them with the neutralized Vitrogen Collagen solution prior to gelation.
Preparation of Vitrogen Fibrillar Collagen Films for Covering Cell Culture Surfaces
A. Materials are the same as described above for preparing collagen gels.
B. Preparation of collagen films
1. Prepare neutralized, isotonic Vitrogen Collagen solution as described above.
2. Cover surface with this solution to a depth 1-2mm (1-2 ml for a 35mm cell culture dish).
3. Incubate for approximately 60 minutes at 37°C to promote gelation.
4. Leave dish uncovered in laminar flow hood overnight or until dry.
5. Rinse film with sterile H2O in order to remove salts and rehydrate film.
6. Film can be used immediately for cell culture or allowed to dry again and be stored for future use.
Preparation of Vitrogen Monomeric Coatings for Covering Cell Culture Surfaces
A. Materials are the same as described above for preparing collagen gels.
B. Preparation of collagen films
1. Place a thin layer of Vitrogen Collagen inside the dish or well to be coated. Vitrogen Collagen may be diluted with 0.01N HCI or 0.05M acetic acid prior to coating, if a thinner layer is desired.
2. Take to dryness in a stream of sterile air or alternatively leave the dish uncovered in a laminar flow hood overnight to allow for normal evaporation.
3. Rinse dish with sterile buffered isotonic saline solution or media to remove residual acid and rehydrate collagen prior to use.
4. Collagen coatings prepared in this manner are nonfibrillar in nature and thus can be distinguished from the fibrillar collagen preparations described above.
REFERENCES
1. Elsdale T, Bard J. Collagen substrate for studies on cell behavior. J. Cell Biol. 54:626-637, 1972.
2. Bell E, Ivarsson B, Merrit C. Production of a tissue-like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential "In Vitro". Proc. Natl. Acad. Sci. 76:1274-1278, 1979.
3. Emerman JT, Pitelka DR. Maintenance and induction of morphological differentiation in disassociated mammary epithelium cells on floating collagen membranes. In Vitro. 13:316-328, 1977.
4. Kleinman H, McGoodwin EB, Rennard SI, Martin GR. Preparation of collagen substrates for cell attachment: effect of collagen concentration and phosphate buffer. Anal. Biochem. 94:308-313, 1979.
5. Bornstein M. Rat-tail collagen as a substrate. Lab. Investig. 7:134-137, 1958.
Collagen gel containing 3T3 fibroblasts (dermal equivalent for raft culture)
Ingredients for 6 x collagen matrices in a 6-well plate;
Roughly 3x106 J2-3T3s (a fully confluent T75?)
1.5ml 10x reconstitution buffer
1.5ml 10x DMEM
12ml rat tail type 1 collagen (>3.8mg/ml)
10N NaOH
Glacial acetic acid (in case)
Method
1. Pre-chill pipettes, keep collagen on ice, as it solidifies above 8oC
2. Mix 1.5ml of 10x DMEM with 1.5ml of 10x reconstitution buffer, keep on ice.
3. Count J2-3T3s and pellet required number 15ml conical tube.
4. Add the 3ml of [1:1, 10x DMEM and 10x reconstitution buffer] and swirl to resuspend the cells, keep on ice
5. Using chilled pipette, add the 12ml of collagen gently to the cells and tilt to mix, avoiding bubbles as much as possible.
6. Add 10N NaOH to bring the pH up to 7. (Judge the pH visually by the phenol red in the DMEM, or use pH paper. Approx 30-60μl will be necessary. Don’t go too far & use glacial HOAc if you have to, but the more mixing the more bubbles.)
7. Pipette 2 2.5ml into each well and incubate O/N
8. In the morning, add 2ml raft media on top of each matrix.
9. Use within 1 week, change media every 2 days.
Buffers
10x DMEM:
Dissolve DMEM powder into 0.1 volume of H2O.
Filter sterilize and store at 20oC in working aliquots. (It may look yellow and not dissolve completely).
10x reconstitution buffer:
Dissolve 2.2g sodium bicarbonate and 4.8g HEPES in 100ml H2O.
Filter sterilise and store at 20oC in working aliquots.
Suppliers
Collagen: Collaborative Biomedical Products (part of Becton Dickinson) Cat # 354236
(It comes in vials of 100mg dissolved in 0.02N HCl at roughly 4mg/ml (ie 25ml). Try to get a batch of at least 3.8mg/ml. If it is stronger than 4mg/ml then dilute it to 4mg/ml with 0.02N acetic acid.)
5 Immunofluorescence & Microscopy
Coverslip Preparations
Clean The Bare Minimum
1) Pick up a single coverslip with forceps and dip into a small (~50ml) beaker containing fresh 95% EtOH. Swirl back & forth a few times & remove.
2) Remove the coverslip from the EtOH and carefully hold it to the flame of a Bunsen burner and let the EtOH burn off. BE SURE that all the EtOH, including any remaining between the forcep blades and the coverslip, has burned off.
3) Hold the coverslip while it cools (~5-10”) then place into appropriate vessel (e.g. one well of a six-well plate).
Acid Washed (it’s not just for jeans anymore)
1) Separate coverslips from one another and place them into a glass container (low-profile screw-cap jar with gasketed lid (preferred) or 500ml beaker) with a lid containing the amount of ddH2O you''ll need to add in order to make a 1M HCl solution. It is important that the cover slips are not sticking to each other because you will need to use one at a time and it''s easier to do this now rather than later. And be careful when changing solutions not to pour out cover slips into the sink which can block the drain.
2) Heat cover slips in a loosely covered glass beaker in 1M HCl at 50-60oC for 4-16h, then cool to room temperature.
3) Rinse out 1M HCl with ddH2O.
4) Fill container with ddH2O and soak for 30min.
5) Rinse container twice with ddH2O.
6) Fill container with 50% EtOH and 50% ddH2O and soak for 30 min, then drain.
7) Fill container with 70% EtOH and 30% ddH2O and soak for 30 min, then drain.
8) Fill container with 95% EtOH and soak for 30 min, then drain.
9) Fill container with 95% EtOH and keep well-covered in appropriate spot (e.g. flammables cabinet).
For Cryin’ Out Loud
Follow the above protocol, but replace the word ‘soak’ with the phrase ‘sonicate in water bath’.
General Immunofluorescent Staining Formaldehyde Fix 11/17/02
1. Prepare all working solutions just before use.
&nb, sp; Fixative (12mls) Permeant (12mls)
1.2mls 10X PBS 1.2mls 10X PBS
1.2mls stock formaldehyde (final=3.7%) 0.3mls 20% Triton X100 (final=0.5%)
9.6mls H2O 10.5mls H2O
2. Remove medium from cells on coverslips and add appropriate amount of fixative (e.g. 1ml/well of a 6-well plate). Fix for 10min at RT without agitation (this is necessary to preserve fine structures). Note: we find it wholly unnecessary to wash with PBS before fixation & find that there is indeed some benefit to avoiding this step.
3. Remove fixative and wash once with 1X PBS.
4. Add permeant to cells and incubate for 10min at RT.
5. Wash once with PBS and block with 1.5% BSA in PBS either overnight at 4oC or for 1hr at RT. Alternatively, you can block in normal serum (same species at primary).
6. Prepare humidified chamber by placing a circle of Whatman 3mm in a 15cm Petri dish, wetting completely with water (drain off excess) and overlaying with a square of Parafilm.
7. In a humidified chamber, incubate cells with primary antibody (diluted in 1.5% BSA in PBS) for 1hr at RT, or overnight at 4oC.
8. Wash with PBS four times, 5min each, with gentle agitation. Alternatively, you can wash coverslips by ‘dunking’ 10 times into 4 successive 250ml beakers filled with PBS, but dunk gently.
9. In a humidified chamber, incubate cells with secondary antibody (diluted in 2% BSA in PBS) for 1hr at RT. If co-staining with phalloidin (e.g. Alexa647 phalloidin at 1:100), include with secondary rather than primary antibody.
10. Wash as in Step 8.
11. Before mounting rinse cells briefly (or dunk 5 times) in deionized H2O to remove salts. Drain by holding edge against a Kimwipe for 2-3sec, then invert coverslip onto a drop of mountant (e.g Permount).
Phalloidin Staining 6-19-2002
General Considerations: The binding of phallotoxins (phalloidin, phallacidin) to fixed actin filaments requires the filaments to be as near their ‘natural’ conformation as possible. Thus, fixation with methanol or acetone, both of which dehydrate samples and alter filament morphology, is contraindicated when staining with phalloidin. Also, the binding of phalloidin to most vertebrate forms of F-actin is very strong and very specific. The incubation times and concentrations suggested below may have to be tweaked, empirically, for optimal staining of a given cell type under given growth conditions. Finally, if combining phalloidin staining with immunostaining, you may opt to include phalloidin in the secondary antibody dilution (at ~ ? the concentration given below), or you may stain with phalloidin (as below) after washing after your secondary antibody.
The protocol below assumes that your cells have been plated onto coverslips (or their equivalent) and your experimental manipulations are complete.
1. Fix cells with 1-3.7% formaldehyde, diluted in phosphate-buffered saline (PBS), for 10-12min at room temperature. (Note: washing cells with cold PBS prior to fixation is not necessary, and in some cases it can actually lower the stability of some finer, peripheral actin structures.)
2. Remove fixative, rinse samples once with PBS, then permeabilize with 0.5% Triton X-100 in PBS, for 5min at RT. (Note: if necessary, lower detergent concentrations, e.g. 0.1-0.2%, may be used).
3. Remove permeabilization solution and wash samples twice with PBS, 3-5min each wash. During washes or blocking step (below), prepare a humidified staining chamber (I typically use a 15cm Petri dish containing a round of Whatman filter paper with a square of Parafilm on top of that. Moisten the Whatman paper with water, drain the excess, and press the Parafilm flatly and evenly on top of the wetted paper. Use this until it literally falls apart.)
4. Incubate the coverslips in PBS containing 1% bovine serum albumin (PBS+BSA) for 15min at RT. (Note: Some labs omit the blocking step and get fine results. I find it helps prevent ‘shroudy’ staining. Also, I most often use phalloidin in conjunction with immunostaining, in which cells are blocked in PBS+BSA).
5. Drain the coverslips by briefly & carefully holding the edge to a Kimwipe, then invert the coverslip onto a drop (30-40μl is more than sufficient for a 22x22mm coverslip) of fluorescent phalloidin diluted in PBS+BSA. Typical dilutions or concentrations are as follows:
Alexa-conjugated phalloidins 1:100 of a 0.2U/μl stock
FITC/TRITC-conjugated phalloidin 100 ng/ml - 1 μg/ml
(Note: Molecular Probes is odd in that they sell you a number of ‘units’, rather than a μg amount, of product. We dissolve 1 vial (300U) in 1.5ml methanol, and use a 1:100 dilution of this. Most other companies sell a μg amount, and so the weight/volume concentrations are used for these.)
6. Incubate at RT for 20min. Avoid prolonged incubations, as background can accumulate.
7. Wash coverslips 2-3 times with PBS, 3-5 min each wash, then rinse once with water. (I usually replace the coverslips in the dish they came from for washing. You may also use the ‘sequential dip’ method, dipping each coverslip 10 times in each of three 250ml beakers full of PBS, then 5 times in a beaker filled with water)
8. Drain the coverslips against a Kimwipe, then mount on slides using whatever mounting solution you’re used to.
A few hints:
- Do not shake or otherwise agitate the coverslips during fixation and permeabilization. This will help preserve fine, peripheral structures.
- Do not let the coverslips dry out at any time during the procedure. Be especially mindful of this when draining coverslips against Kimwipes & when transferring coverslips into & out of their original plate.
- Many labs report fine results without using BSA to block, and also with less extensive washing before mounting. Follow their lead at your own risk.
Fluorescent Staining Actin Cytoskeleton and Nuclei 06/01/03
Of course, these targets may be stained separately. They are offered together here because the protocols for both are very short & quick. Three quick usage notes. 1) We routinely dilute the Molecular Probes Alexa-phalloidins according to their instructions (i.e. dissolve 300 units in 1.5mls of methanol to give 200 units/ml). Now, I don’t know what a ‘unit’ of phalloidin is, but this stock is effectively 100X. 2) For DAPI (D-1306 from Molecular Probes), we dilute the supplied 10mg in 9.5mls of H2O, giving a stock of 3mM. This is ~5x less concentrated than the manufacturer’s protocol, but solubilization is not a problem and it still gives a 5000-10,000X stock solution (final working conc = 300-600nM). 3) If staining only with actin or DAPI (i.e. if no immunostaining is involved), blocking is unnecessary and washing is largely optional, although a dunk or two in PBS might help lower the background.
Reagents
Alexa-Phalloidin (100X stock) 300 U into 1.5mls MeOH
DAPI (3mM stock) 10mg into 9.5mls H2O
(Use at 1:5000 to 1:10,000)
1. Fix cells using standard formaldehyde fix or the glutaraldehyde/formaldehyde fix for microtubules. DO NOT use methanol fixation for subsequent staining of F-actin with phalloidin.
2. Incubate the cells with Alexa-conjugated phalloidin (diluted 1:100 in PBS or PBS/BSA) for 30min or with DAPI (diluted 1:5000-1:10,000 in PBS or PBS/BSA) for 5-10min at RT.
3. Wash cells briefly with PBS and rinse or dip once in H2O before mounting.
4. If using phalloidin as a counterstain during immunofluorescence, include it at a dilution of 1:100 in the secondary antibody solution and stain as usual. For counterstaining with DAPI, either do a third, short incubation with DAPI at 1:5000 - 1:10,000 after the excess secondary antibodies have been washed off (preferred) or include the DAPI at 1:25,000 in the secondary and stain as usual.
Microtubule Fixation / Immunofluorescence 11/25/02
While microtubules can also be readily visualized in MeOH-fixed cells, that fixation method alters or wipes out phalloidin-based staining of the actin cytoskeleton. The following protocol uses a mixture of formaldehyde and glutaraldehyde, and affords excellent staining of microtubules, F-actin, and almost every other antigen that is typically visualized after formaldehyde fixation.
Buffers
2X CB 20mM MES pH 6.2 2X Sucrose 22.2% in H2O
280 mM NaCl
5mM EGTA (Store both 2X CB and 2X sucrose at 4oC, and be mindful
10mM MgCl2 of yuck growing in sucrose over time)
Fixative 5mls 2X CB
1ml formaldehyde (stock = 37%; final [ ] = 3.7%)
200 μl glutaraldehyde (from 25% EM-grade stock; final [ ] = 0.5%)
125 μl 20% Triton X100 stock (final [ ] = 0.25%)
2.5ml 2X sucrose
1.18ml H2O
Quench 0.5mg/ml sodium borohydride in 1X CB (~2 spatula tips full/10mls is fine)
(make fresh during fixation; solution bubbles vigorously no sweat)
1. Remove media from cells and gently add fixative. Fix at RT for 15min without agitation (this is necessary to preserve fine structures). Note: it is unnecessary to wash with PBS before fixation & there may be some benefit in avoiding this step.
2. Remove fixative and gently add quench to cells. Be mindful of floating coverslips. Quench at RT for 8min. Gently flick samples once or twice during this period to dislodge bubbles.
3. Wash cells once with PBS and block in PBS + 2% BSA, overnight at 4oC or for 1hr at RT.
4. For microtubule staining, use the DM1A anti-a-tubulin monoclonal (from Sigma), at 1:500 to 1:1000 for 1hr at RT. This antibody is very strong and very clean. Counterstain with other antibodies as desired.
5. Wash once with PBS-0.1% Tween20 and three times with PBS, 5min each, with gentle agitation. Alternatively, you can wash coverslips by ‘dunking’ 10 times into 4 successive 250ml beakers filled with PBST and PBS, but dunk gently.
6. Stain with appropriate secondaries and phalloidin as desired (e.g. Alexa594-anti-mouse at 1:1000 and Alexa488-phalloidin at 1:100).
7. Wash as in Step 5.
8. Rinse coverslips briefly with water (i.e. dunk 4-5 times in a beaker of water) before mounting.
Immunofluorescent Staining of Pseudopod Preps 2/17/03
The following protocol assumes that cells have already been plated on filters and stimulated to send protrusive elements (pseudopods) through the filter pores. If you ultimately wish to image the entire prep in one series (i.e. take a Z-stack from the bottom of the pseudopodia up through the filter and through the cell body), use polyester filters (e.g. CoStar Transwell Clear) rather than the more opaque polycarbonate. Although certainly not totally ‘clear’, the polyester membrane will allow a reasonable amount of light through for imaging.
1. After pseudopodia have formed, transfer the Transwell filter inserts into a separate multiwell dish containing your fixative of choice[4] - for example, 3.7% formaldehyde in PBS. Add an appropriate amount of fixative to the top chamber of the insert. We find it wholly unnecessary to wash with PBS before fixation & find that there is indeed some benefit to avoiding this step.
2. Fix for 10min at RT without agitation. This is necessary to preserve fine structures.
3. Remove fixative from top & bottom chambers and wash once with PBS (by adding an appropriate amount to top and bottom chambers).
4. Permeabilize with 0.5% TritonX100 in PBS for 10min at RT.
5. Wash once with PBS and block filters, still in the inserts, overnight[5] at 4oC in 12 % BSA in PBS (again, by adding an appropriate amount to top and bottom chambers). The exact concentration seems more to do with the cell type, target antigen, and antibody quality than with the nature of the pseudopod prep.
6. Excise filters with a scalpel (e.g. #10 blade), taking extreme care not to let the filter fall against the side of the insert (or fall altogether). The last few millimeters will be difficult because of lack of counter-tension. If too problematic, cut to this point then gently tear the rest of the filter from the insert with forceps, grasping near the remaining margin of contact. Do not attempt to image this area.
7. Minding their ‘sidedness’, place filters gingerly on a drop of diluted primary antibody solution[6] and cover the topside of the filter with another, equal drop of solution. Stain for at least 2hrs at RT, or overnight2 at 4oC.
8. Wash filters by removing from antibody solution and placing in an appropriate multiwell dish containing a sufficient volume of PBS to prevent filter from immediately settling to the dish bottom (e.g. 4mls in each well of a 6-well dish). Place dish on shaking platform set fast enough to keep filters off the bottom but submerged. For multiple samples, work quickly to prevent filters from settling[7]. Shake for 10-15min.
9. Wash filters twice more for 10-15min each by removing to a new multiwell dish with fresh PBS. When removing filters from wells, be very careful not to scrape the filter against the side of the well.
10. Incubate filters in diluted secondary antibody solution as in Step 7. If co-staining with phalloidin, include with secondary rather than primary antibody.
11. Wash filters three times for 10-15min each as in Steps 8 & 9.
12. For mounting, place a drop of mountant (your favorite we use Permount, glycerol, or even microscope immersion oil) on a long rectangular (25x60mm) coverslip. Gingerly lay filter on drop, then add another drop of mountant on top of filter[8]. Top with another coverslip (standard square or round will do) and, if desired, seal edges with clear nail polish, taking care that no polish comes in contact with filter.
DAPI Staining Transwell Inserts (Migration Assay) 11/01/2002
1. Coat inserts with ECM protein (e.g. Col I @ 10-20μg/ml, Fn @ 25μg/ml) by placing insert on a drop (~30μl) of diluted ECM solution, then adding an equal volume of the ECM solution to the inner chamber of the insert. Coat at RT for 2hrs or O/N at 4oC.
2. Place inserts back into plates & wash 2x with PBS.
3. Seed an appropriate number of cells in an appropriate volume (e.g. 2x104 in 0.5ml for a 12mm insert) into the top chamber of the insert. Add serum-free medium to the bottom chamber (e.g. 1.5ml for a 12mm insert / 12-well plate).
4. Let the cells adhere & spread for 4hrs at 37oC.
5. Replace the medium in the bottom chamber with serum-containing growth medium and incubate at 37oC for 16-24hrs to allow migration to occur.
6. Fix cells in 20oC methanol for 20min, then bring to RT by replacing cold MeOH with RT PBS.
7. Remove unmigrated cells from the inner chamber/top of the membrane with a cotton swab. Steps 6 and 7 can be reversed, but I prefer this order.
8. Stain the cells on the bottom of the membrane by placing the insert on a drop of DAPI, diluted to 300nM in PBS, and incubating at RT for 5min. Washing is unneceassry, but will not damage the sample.
9. Excise the membranes and mount, minding their sidedness, between glass slides and coverslips using an appropriate mountant (e.g. Permount).
10. Capture > 3 fields (@ < 400X magnification) per insert using UV filter on fluorescent microscope.
11. For easy analysis of high-count fields, use ImageJ as follows:
a) Open image in ImageJ. ‘Image’à ‘Type’à ‘8-bit’ to convert to 8-bit grayscale image (if not already in this format).
b) ‘Binarize’ or threshold the image (make it B&W) by either of two methods:
i) ‘Process’à ‘Binary’à ‘Threshold’ (a very automated method)
ii) ‘Image’à ‘Adjust’à ‘Threshold’; use slider to adjust threshold (red areas will become black in the binary image.
c) ‘Analyze’à ‘Analyze Particles’; Adjust Min/Max sizes (empirically), set ‘Show’ to ‘Outlines’, check boxes for Display Results, Exclude Edge Particles (if desired), Clear Results Table, and Summarize. Click ‘OK’.
‘Inverse’ Transwell Plating (for plating on insert bottoms) 10/31/2003
From:
Dan Focht
Bioptechs
3560 Beck Road
Butler, PA 16002
dan@bioptechs.com
www.bioptechs.com
We plate our cells on the basal surface of the transwell insert in the following manner.
1. A bio-compatible piece of tubing is cut into a small cylinder of suitable geometry as to provide a shallow well and fluid barrier when placed over the distal end of the insert.
2. A plug made of silicon is placed into the tubular portion of the insert to prevent leakage through the membrane.
3. The insert is inverted (membrane side up) and cells are poured into the well defined by the cylindrical tubing surrounding over the membrane.
4. The cells are returned to the incubator and allowed to plate in this inverted orientation for 45 minutes.
5. The plug and tubing are removed from the insert and the insert is returned to the tray with appropriate media and allowed to divide until confluent. We have had much success with this plating technique.
6. When the cells are confluent the insert is placed into a Bioptechs Delta T4 Transwell Adapter which is then lowered into a Delta T dish on the microscope. The cells are now facing down for easy imaging through the coverglass bottomed self-heating Delta T dish. See Bioptechs web site for details on equipment and contact Bioptechs directly for additional information regarding membrane insert micro-observation accessories.
7. Cells on the membrane can be perfused on either the apical or basal surface during microscopy if necessary.
6 Appendix
Some Common Solutions
10x PBS
20.45 g NaCl
0.465 g KCl
10.142 g Na2HPO-4 * 7 H2O
0.545 g KH2PO-4
Add distilled water to 250mls, stir to dissolve
Homemade Coverslip Mounting Media
20mM Tris pH 8.0
0.5% N-propyl gallate
90% Glycerol
Store at 4oC
Valap
(from Waterman-Storer, C.M. Microtubule/organelle motility assays. In: Current Protocols in Cell Biology, J.S. Bonifacino, M. Dasso, J.B. Harford, J. Lippincott-Schwartz, and K.M. Yamada, eds. John Wiley, NY.)
Put 50 g Vaseline, 50 g lanolin, and 50 g paraffin (all from Fisher) in a 1 L Pyrex beaker
Heat on "low" on a hotplate, stirring occasionally, until all components are melted & well mixed
Pour into several small screw-cap jars (~50 ml capacity)
Store at RT
PBS
8.18 g NaCl (140 mM NaCl)
0.186 g KCl (2.5 mM KCl)
0.218 g KH2PO-4 (1.6 mM KH2PO-4)
2.15 g Na2HPO-4 (15 mM Na2HPO-4)
distilled water to 1 liter
Store at RT
1M MgSO4
24.074 g MgSO4
distilled water to 200 ml
store at RT
1 M HEPES pH 7.0
119.15 g HEPES (free acid)
distilled water to 400 ml
add solid NaOH a few pellets at a time while mixing until the pH is ~6.8
add concentrated NaOH dropwise until pH = 7.0
distilled water to 500 ml
sterile filter (do not autoclave) and store at 4oC
1 M PIPES, pH 6.9
151.2 g PIPES (free acid)
distilled water to 400 ml
add solid NaOH a few pellets at a time while mixing until the pH = ~6.7
add concentrated NaOH dropwise to until pH = 6.9
distilled water to 500 ml
sterile filter and store at 4oC
0.5 M EDTA
16.81 g EDTA (Sodium Salt)
distilled water to 90 ml
Adjust pH to 7.0
Distilled water to 100 ml
0.5 M EGTA Stock
19.02 g EGTA (Sodium Salt)
distilled water to 90 ml
Adjust pH to 7.0
Distilled water to 100 ml
1M dithiothreitol (DTT)
1.542 g dithiothreitol
distilled water to 10 ml
disburse into 500 μl aliquots and store at -20oC
10 M NaOH Stock
40 g NaOH
distilled water to 100 ml
5 M NaCl
292.2 g NaCl
distilled water to 1 liter
LB Broth (Lennox formulation)
10 g Bacto-tryptone
5 g Bacto-yeast Extract
5 g NaCl
Distilled water to 1 liter
Stir until dissolved.
For Miller formulation, use 10g NaCl
