Z3221319 Lab Protocols For This Project

Relevant Lab Protocols For Cell Culture, Immunohistochemistry and Molecular Bioogy)

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Protocols will be added as they are learnt and include cell culture, molecular biology and immunohistochemistry.

Using the Biohood

1. Turn on (hold down 'on' button until beep sounds).
2. Sterilise with UV light for 20 minutes (hold down UV button until lights turn on - timer is automatically set to 1 hour)
3. Raise glass sheath (hold down 'up' button - Alarm will sound until air velocity is stabilised, therefore wait until red air flow light turns green).
4. Wipe down bench with 1% Bleach and Kim Wipes
5. Wipe down bench with 70-80% Ethanol spray (ANY item entering the hood should be sprayed and wiped with ethanol - including the gloves you wear.)
6. Keep cleanest materials at the back, working materials (more likely to get contaminated) at the front. Keep middle area clear for air flow.

Preparing 1 Litre of DMEM (Dulbecco's Modified Eagle's Medium) Stock Solution

Instructions are on the packaging for the DMEM powder sachets. Most of this work is done in the G16 lab but filtering is in the PC2 lab. These instructions indicate how to convert low glucose powder mix (1000mg/L) to high glucose mix (4500mg/L). The powder used was supplied by Invitrogen.

NB: The steps to create calcium/magnesium free phosphate buffered saline (PBS) from a powder, are the same except ignoring steps 4 and 5

1. Measure out 5% less distilled water than desired into volumetric container closest to desired volume. i.e. 950mL.
2. Add powder and stir gently using magnetic plate and stir rod/pellet
3. Rinse out inside of packaging to empty powder remnants into flask
4. Add 3-7g of NaHCO3/L (if it's a bicarbonate-free powder).
5. Add 3.5g of D-Glucose. (This is only when converting from low to high glucose powder mix).
6. Stir to dissolve
7. Dilute to desired volume with distilled water.
8. Adjust pH to 0.2-0.3 below desired final pH using 1M NaOH or 1M HCl - checking pH using pH meter. i.e. pH should be made to 7.1-7.2
9. Close container with parafilm
10. Sterilise by membrane filtration

Membrane Filtration of DMEM Stock Solution

This describes the process for a reusable filter, though the single use filters are similarly designed.

1. Spray and wipe filter and 2x500ml tissue culture flasks, with ethanol. Filter should be autoclaved.
2. Ensure pump is connected to waste and overflow flasks
3. Screw sterile filter onto autoclaved flask (ensure you do not touch pump attachment or point of screw-on).
4. Tighten seal on filter by screwing white ring clockwise
5. Attach pump tube to filter
6. Remove filter cap and pour in 500mL
7. Turn on pump, wait for 500mL to dribble through
8. When done, remove pump by first pinching it near the attachment, to prevent any air sucked into the filter being taken from the tube (the pump creates a negative pressure). Instead the air will be sucked from the sterile biohood air. After tube is removed, then turn off pump.
9. Repeat process for next flask.
10. When done, open seal on filter (anticlockwise), and check the filter is intact (should see visible ring indicating a seal). If filter is intact, throw away. If it is not intact, must refilter the solution.
11. Label flasks. E.g. 'DMEM High Gluc. Initials. Date.
12. Place them in fridge.
13. Rinse filter and dry in oven (autoclaving not necessary for DMEM)
14. When dry, assemble new filter and autoclave.

Double Checking for Sterility of DMEM Stock Solution

1. For the two 500mL flasks of stock solution created above, get 2 UV irradiated culture plates, spray clean whole packet (if new, unopened packet), before opening in the hood and removing 2 plates.
2. Label plates. E.g. DMEM Sterility Test. 1 or 2. Initials. Date
3. Get two single use 1mL pipettes (plastic with bulb at top). Plate out 1mL of each stock solution into relevant plate (ensure stock solutions are numbered 1 or 2 as well.
4. Leave plates in incubator overnight and check for sterility the next day. This is done with an inverted phase microscope - see below.

Using an Inverted Phase Microscope

NB the x4,10,20 and 40 magnifications below the target (light shines from above).

NB the plate carrier is for holding of 24 plate wells. An adaptor can be placed into the plate area for round plates.

1. Calibration: Turn light on and remove one eye piece. In its place, insert the 'CT' telescope.
2. There are three phase rings (in the phase filter) for the x4, x10/20 and x40 magnifications. These rings can be slotted into the top part of the microscope near the light source and should be changed as appropriate, each time the magnification is changed. The main one used in cell culture is the x10/20. Calibration should be undertaken for each magnification using the separate calibration slide (located in the top draw on the left of the desk).
3. When looking down the telescope at the calibration rings, the offset grey ring should be moved until it fits over the ring of light. This is done by physically moving the phase ring apparatus that was slotted in earlier.
4. After Calibration: Replace eye piece and adjust distance between eye pieces as appropriate.
5. Place culture plate under light and focus microscope.

NB: Can also use green filter (slots into top part of microscope near light source). This often makes it easier to distinguish cells.

Converting DMEM Stock Solution into DMEM f10 P/S (i.e. Addition of 10% fetal calf serum and also penicillin/streptomyocin)

NB: for a P/S with penicillin 5000 units/mL, to make 50ml of DMEM f10 P/S, we will add:

• 45mL DMEM stock solution
• 5mL FCS
• 0.5mL P/S

NB: for the PC12 cells, must create DMEM f5 h(horse serum)10 P/S. We will add:

• 42 mL DMEM stock solution
• 2.5mL FCS
• 5mL HS
• 0.5mL P/S

1. Set up water bath at 37 degrees celsius, to thaw frozen vials of fetal calf serum (fcs) and penicillin/streptomyocin (P/S). When thawing, only raise water level to just below the caps/seal of the vials. The seals should not get wet.
2. Take a 10mL plastic pipette, open packet partially and insert into the pipette man. Then draw pipette out of packeting. (On the pipette man, the top button draws solution up into the pipette and the bottom button dispenses).
3. Pipette 45mL DMEM stock into two 50mL tubes (We are creating 2x50mL aliquots). NB. 45mL = 4 full pipettes from the 10mL pipette and one half.
4. Mix thawed FCS vial by inverting, draw up about 10mL (because the previous pipette has only been in DMEM, it is not necessary to change it. When not using the pipette, rest it sitting on the pipette man, so that the plastic part is in the air and not touching anything).
5. Place 5mL in each 50mL tube
6. Take a 1mL plastic pipette to be attached to the pipette man (alternatively can use pipettes with tips), and add 0.5mL of P/S to each 50mL tube
7. When done, refreeze FCS and P/S.

Plating out DMEM f10 P/S or DMEM f5 h10 P/S for reception of cells

NB: As we were going to plate out 3 cell types - RGC-5, B35, PC12 - we used 6 x 10cm plates

1. When opening plate packet, ensure you spray first and take care not to open any plates except the one you're using.
2. Pipette 8mL of DMEM f10 P/S or DMEM f5 h10 P/S into each plate as necessary.

Collecting the cells

NB: Whenever transporting cells between rooms, always double contain in a bullet rack (test tube rack) and esky. The cells are stored in liquid nitrogen in G14 and the PC2 lab. Each liquid nitrogen dewer contains a number of canisters, which then each contain numerous 'canes' (racks of vials). Whenever removing a vial, it MUST be logged in the log book by the dewer or in your lab book. This includes recording the contents of the tube, tube number, date originally frozen and by whom. If the contents are already recorded in a lab/log book, when removing this tube, ensure you cross out that tube in the book, initial and date it.

In this case, we removed 3 vials from the dewer in the PC2 lab:

• PC12T frozen by MV 22/01/98, Tube 7
• B35 frozen by MH 3/10/98, Tube 19
• RGC5 frozen by MS 29/11/99, Tube 5

NB: Face mask and blue cryo gloves when handling liquid nitrogen.

1. Cells need to be thawed as rapidly as possible. Do this by holding the tubes (one at a time) in a water bath (remember that water should not touch the cap/seal). DON'T be rough with the tubes - the cells are old and fragile. Move slowly, and don't mix/invert.
2. Thaw takes a few minutes, when all the ice is gone.
3. Spray and wipe vial.
4. Get 3 single use plastic pipettes.
5. Using a single use plastic pipette, draw up 1mL slowly, and drop 0.5mL around the first DMEM f10 P/S plate (from the 6 that you created previously). Drop the other 0.5mL around the second plate.
6. Label these plates (initials, date, cell line, plate #) and put them aside.
7. Repeat steps 1-6 for each cell vial (i.e. 3 in total - see above).
8. Once all plates have cells in them, check one plate from each pair under the microscope for cell viability and density. The density should be roughly estimated and recorded along with the date and time and magnification of the microscope.
9. Any remaining DMEM f10 P/S or f5 h10 P/S goes into fridge.

NB: In this case, the recorded densities at 10x magnification at 1:40pm on the 24/06/09 were:

• RGC-5 ~ 100 cells
• B35 ~ 100 cells
• PC12 ~ 70-80 cells with clumps (not counted).

Changing the media for plated cells

NB: This is generally done twice a week, but for cells that have just been thawed (as above), it is a good idea to change it the next day.

NB: Always check cells with microscopy prior to media replacement. If cell debris is rife, wash cells with PBS or HBSS (mostly the latter), before replacing media.

1. Pre-warm DMEM f10 P/S or f5 h10 P/S in water bath, spray and wipe before entering hood.
2. Take glass pipette out of metal canister in biohood, and attach it to the pump (ensuring you don't touch the tip).
3. Turn pump on
4. Put two plates to be replaced, side by side with lids off.
5. Take plate by its side and tip on its edge so that all the media pools in a corner.
6. Place pump into pooled media (right into corner of plate) and draw out media - don't need to 'suck it dry'.
7. Take pipette man and 10mL plastic pipette, to transfer 8mL of new, pre-warmed DMEM f10 P/S media to plate.
8. After using pump, clean line by attaching one of the used glass pipettes (from the metal canister), sucking up about 1mL of bleach and then about 1mL of ethanol.

Using phase microscope to take photos of cells (using SPOT QUANT software)

1. At each magnification, set the white balance by pressing the sidebar button looking like an inverted tricolour flag on a white piece of paper. Nothing should be in the target area when you do this.
2. To set calibration for each magnification, take out phase filter, put in calibration slide (top draw on left) and at 4x focus on the 1mm long scale. To see the image on the computer screen, click the 'live' button on the sidebar. When it's appropriately focussed (often the screen focus is different to what is seen down the microscope. There is also some delay between microscope adjustments and adjustments on screen), click the 'get image' button (below the 'live' button) on the sidebar, to take a photo of it.
3. The photo will be in a window behind the 'live' window. So close the 'live' window and click on the photo. Then click file >> save as - to save it in a folder.
4. To calibrate for this magnification (4x), click setup >> calibration >> add. Type in a name such as '4x' and drag a line across the 1mm (the smallest unit of measurement on the scale is 10 microns). Then adjust the length value in the caibration box and click 'add'.
5. Repeat steps 2 and 3 for each magnification.
6. Taking photos of cells involves the same procedure, except it is a good idea to replace the phase filter and green filter.

NB: When looking at the cells under the microscope; round, phase bright (with a halo like effect) cells are pre-division cells. Dead cells are floating, with an irregular shape and no phase brightness. Growing cells may appear as fibroblast-like or may have processes called neurites (once they are differentiated they become known as axons or dendrites).

NB: To look for bacterial infection - should see black spots about a 10th the size of the cells. Fungal infection is visible by filamentous strands.

NB: As well as the focus being different on screen compared to the microscope, the pictures taken are also often lighter/brighter/overexposed compared to the microscope image. Keep this in mind.

NB: When taking photos of cells, do not leave cells out of incubator for more than a few minutes.

NB: A record of photos taken are on a separate page accessed from the project home page.

Splitting Confluent Plated Cells and Cell Quantification with a haemocytometer

NB: Must first detach cells from plastic plates using calcium/magnesium free PBS (it deprives the cells of calcium required to maintain intercellular adhesion) and trypsin/EDTA solution (digests intercellular adhesion proteins).

1. Aspirate out growth media from plate, using pump/glass pipette as usual
2. Add 10mL calcium/magnesium free PBS using pipette man, and gently swirl.
3. Aspirate again
4. Using a 1mL pipette man, add 0.5mLtrypsin/EDTA solution drop by drop around the dish. Swirl gently.
5. Incubate at 37 degrees celsius for 1 minute, then tap plate moderately on sides to dislodge any stuck cells. Check under microscope. NB: Under the microscope, the cells that have dislodged, no longer exhibit a flattened appearance, but rather revert to their original round appearance. If there are still too many attached/still confluent, incubate for another minute and check.
6. Add 5mL growth media (using 1mL pipette) and tilt plate to make sure all cells are rinsed down into media.
7. Tilt plate and pipette cell suspension into a 10mL growth tube.
8. Centrifuge tube at 1000 revs for 5 minutes (remember to balance). When centrifuge is complete, remove whole centrifuge jars, wipe with alcohol and reopen in the hood. Also rewipe the tubes.
9. Slowly aspirate off the media by touching the pipette against the surface/edge of the tube, near the meniscus and rotating around the tube. Leave a little bit of media on top of the cells.
10. Add in 5mL of new growth media.
11. Resuspend the cells using a disposable pipette (draw media up and down without making bubbles, to break up the clumps of cells). NB: Here, because we had so many RGCs, we made a 1:10 dilution by transferring 100 microlitres cell suspension into a yellow sample tube, and adding 900 microlitres of DMEM f10 to it.
12. At this stage we set up the haemocytometer by placing a spot of water on the two glass bars where the coverslip will attach to the haemocytometer slide. The coverslip should cover both the visible crosses.
13. Using a disposable pipette, a drop of cells from step 11, is placed at the edge of the coverslip and capillary action pulls it through.
14. Through the microscope, focus on the 1mm x 1mm square that is divided into 0.25 mm x 0.25 mm squares (i.e. divided into 4). Using the cell counter, count all cells in this area, including clumps of cells as 1 cell and counting all cells lying on the right and bottom borders, but not the top and left. Also, the well of the slide is 0.1 mm deep.
15. E.g. from two counts of different 1mm x 1mm squares for the B35's, we averaged 120 cells. Therefore, 120 cell/mm^2. If the well is 0.1 mm deep, then we have 120 cells in 0.1 mm^3. If 1mL = cm^3 = 1000 mm^3, the volume of the well = 0.1mm^3/1000mm^3 = 0.0001cm^3. Therefore, we have 120 cells/0.0001 mL, which is 1,200,000 cells/mL. In the original 5 mL tube, therefore, we had 6 million cells.
16. Replate solution from step 11 into new plates pre-prepared with 8mL growth media. We usually split 1 plate to 5 plates, but we only needed 2 plates. So today we only split a B35 plate and RGC-5 plate into 2 plates each. (The PC12's are still growing and were not confluent at this stage). When replating, add 1mL of cell suspension to each plate. (Just to see the difference under the microscope, we placed 1 ml and 0.5 mL for the B35's and 0.5ml and 0.25mL for the RGC's.
17. Aspirate (using glass pipette and pump) any excess cells/waste to dispose of them.

Preparing glass coverslips in a 24 well plate (including coating with collagen)

1. Glass coverslips should have already been autoclaved in glass petri dishes lined with some paper (makes the slips easier to pick up). Also, should have autoclaved forceps/fine tweezers ready.
2. In the biohood, use the forceps/tweezers to pick up each coverslip and drop it in a well of the 24 well plastic plate (1 slip/well).
3. When all wells full, tap side of plate so that coverslips lie flat.
4. Use a disposable pipette to add enough collagen to cover glass coverslip in each well (no precise volume, just ensure that meniscus of collagen solution doesn't touch coverslip. I.e. the coverslip should be completely covered). NB: That for L-lysine can also be used. Furthermore, for our experiment, we found some old collagen which we refiltered using a syringe filter (see below)
5. Leave wells to soak in collagen for 20-25 minutes
6. Use the same disposable pipette to suck up collagen out of each well (whilst holding the plates at 45 degrees - don't need to suck the wells dry). Place this collagen in a new 50mL tube labelled: 'Collagen used once'. Collagen can be used a number of times before it become significantly altered. NB: Collagen can be stored in the fridge.
7. Leave the remaining plate/wells to dry in the biohood overnight (i.e. with the biohood on and the air flowing through).

Using a filter syringe

For when you need to filter a small amount of solution, quickly - and when it's not worth opening an autoclaved reusable filter.

1. Draw up solution in 10mL syringe.
2. Open and attach 0.2 micrometer syringe filter (squeeze packet by side and twist filter onto syringe to lock on)
3. Apply pressure to syringe to filter solution drop by drop.
4. To draw up more solution, carefully place filter back into packet, and remove from syringe. Draw up more solution in syringe and then repeat proceedure.

Reassembling and autoclaving reusable filters

1. Open used filter and place 0.2 micrometer membrane filter (from white box in third draw under the apple mac in the lab) over the red o-ring (this must be completely covered). Note that the membrane filters are the white pieces in the box (not the blue pieces). The blue pieces ensure that you can handle the filters without touching them as much as possible.
2. To ensure filter sticks down well, drop water over it liberally when it is placed over the o-ring.
3. Reassemble filter (do not tighten white ring too much - leave a little loose).
4. Bag and autoclave.

Making Paraformaldehyde Fixative from Paraformalydehyde powder

1. For the purposes of fixing cells on glass coverslips, the fixative needs to be at 4% weight/volume. Therefore, add 2g paraformaldehyde powder to a 50mL tube. A face mask should be worn when handling the powder. The powder can be found in G16.
2. Add water up to about 25mL - shake tube to try and dissolve as much powder as possible.
3. Add a drop of NaOH from the G16 lab, then shake some more.
4. Place in a beaker in the microwave (in Dr. Hill's CBL) for 5 seconds. DON'T OVERHEAT/BOIL! Then shake. If still not dissolved, give it another 5 seconds.
5. If still not dissolved, add more water to 35mL and microwave for another 5 seconds.
6. When dissolved, add in 5mL PBS 10x (Stock in Dr. Hill's CBL. NOT THE CA2+/MG2+ FREE PBS!).
7. Use pH stick to check pH is roughly 7.4. (The green square on the pH stick is the best indicator). If it's too high, add a drop of HCl from G16.
8. Top up to 50mL with water.

Fixing Coverslips

1. Using one glass pipette for all wells, aspirate media.
2. Using disposable pipette, cover each coverslip with PBS (NOT CA2+/MG2+ FREE PBS) made from PBS 10x stock in Dr. Hill's Lab. Add 5 mL 10x PBS and make up to 50mL with 45mL water.
3. Using glass pipette, aspirate PBS from all wells.
4. Using disposable pipette, add paraformaldehyde to each well (enough to cover coverslip) and leave for 20-30 minutes
5. Using disposable pipette, remove paraformaldehyde from each well and add in some more PBS to cover the coverslips in each well (keep them hydrated).
6. Store paraformaldehyde in fridge with parafilm around it.

Making Differentiation Media

In the 24 well plate Number 2, we used DMEM F5 + 1mM dibutyryl Cyclic AMP (dbcAMP) for all wells.

1. Add 5mL of existing DMEM f10 P/S to 5mL DMEM high gluc. Mix.
2. From this 10mL stock, to make 6mL of differentiation media (250 microlitres in each of the 24 wells) take 5940 microlitres of the stock and add to it 60 microlitres of 100mM dbcAMP (i.e. dilute it 100 times to make it 1mM).

The TRIzol method of extracting RNA from confluent flasks of cells

NB: The procedure followed was that provided by GIBCO (as referenced by Dr. Hill on the ILP home page). Trizol Protocol PDF

1. Aspirate media out of flask
2. Wash flask with 10mL PBS (NOT Ca2+/Mg2+ free) twice and swirl. Aspirate out the PBS each time.
3. Homogenisation - lyse cells by adding 5mL of of TRIzol reagent to each flask. Leave for a few minutes and shake to help lyse cells. (The solution will take on a 'gooey' consistency).
4. Using disposable pipettes, suck out as much solution from the flask as possible and transfer to RNAase-free 10mL tubes (handled ONLY with gloves on). Use the pipettes to mix the tubes.
5. Centrifuge at 1000g for 10 minutes to see whether a pellet forms (cell debris/larger molecules - transfer supernatant into new tubes if this occurs).
6. Place 1mL of the original 5mL solution into each of 5 1.5mL eppendorf tubes.
7. Phase separation - Leave samples at room temperature for 5 minutes and add 0.2mL chloroform per 1mL of TRIzol reagent. Cap and shake tubes vigorously by hand for 15 seconds and leave at room temp for further 2-3 minutes.
8. Centrifuge ar 12000g for 15 minutes at 2-8 degrees celsius (in Dr. Hill's CBL lab).
9. RNA precipitation - following centrifugation, transfer aqueous phase (on top) to fresh tube, save organic phase (lower red phase and interphase)for later DNA and protein extraction. Add 0.5mL isopropyl alcohol to each 1mL TRIzol reagent originally used, to the aqueous phase - this precipitates the RNA (often invisible at this stage).
10. Leave at room temp for 10 miuntes then centrifuge at 12000g for 10 min at 2-8 degrees celsius. The pellet is seen now as a very slight, gell-like, white precipitate on the side of the tube-bottom.
11. Remove supernatant and wash RNA pellet with 75% ethanol (1mL for every 1mL TRIzol reagent originally used). Mix by vortex and centrifuge ar 7500 g for 5 minutes (2-8 deg cel).
12. Redissolving the RNA - Can be done with 0.5% SDS or RNAase free water (see invitrogen website above). We redissolved in 100% formamide (deionised) so that it could be stored at -70 deg cel. We used 50 microlitres of formamide.

The TRIzol method of extracting DNA and Proteins from harvested cells

NB: The methods used here were the same as provided in this protocol - Trizol Protocol PDF. The only changes were due to the fact that our eppendorf tubes that could fit in out cooling centrifuge (2-8 deg cel) could only hold 1.5mL. Therefore for the protein precipitation step (step 1 of protein isolation), I split each tube of phenol-ethanol supernate into two tubes (approx. 385 microlitres in each) so that instead of needing to add 1.5mL of isopropanol to each tube I only had to add 0.75 mL (halved). This meant that instead of having 5 protein samples for each cell-line, we had 10. This method still seemed to produce a considerable pellet following centrifugation at the end of step 1.

For the washing steps, as we could not add 2mL of the guanidine hydrochloride to each tube, we will just give them an extra wash. (This step has not yet been done, as the protien pellets are currently stored suspended in 0.3M guanidine hydrochloride in 95% ethanol - where they may remain for 1 year at -5 - -20 deg. cel.

Rough method of freezing down cells for storage at -80 deg cel.

1. Prepare the freezing media (This should be done anew each time cells are frozen down - i.e the freezing media should not be stored). Note that the overall solution in which the cells are suspended needs to be 10% DMSO. As we add freezing media to normal growth media at a ratio of 1:1, there thus needs to be 20% DMSO in the freezing media. There should also be a minimum of 20% FCS - though this can be higher. I did it at 40%. Therefore, to make 15mL of the freezing media, I first used a synringe to filter 50mL of DMSO. 3mL of this (20%) was added to 6mL FCS (40%) and 6mL DMEM (40%).
2. Split cells as normal to pellet stage (i.e. just after centrifugation)
3. Resuspend pellet (as normal with a disposable pipette) in 2.5mL of normal growth media and 2.5mL of freeing media (5mL total).
4. For each cell line, create 3 ampules (use orange vials for cryogenics) with 1.5mL in each. These should be stored in a foam esky in the -80 deg. fridge.

Immunohistochemistry - staining and mounting coverslips

The coverslips involved are those created previously in 24 well plates (see above). The practice run used two primary Ig's (double staining with anti-beta-actin from mouse and anti-tau from rabbit). The secondary Ig's were FITC goat anti mouse (therefore, B-actin will fluoresce green) and Rhodamine anti Rabbit (tau - red). The coverlips that were used up were 2 undifferentiated and 2 differentiated for both HT4 and RGC-5. And 1 Undifferentiated RGC-5 as a control (only secondary Ig is incubated with the control - to determine background/autofluorescence).

1. Determine primary antibody dilutions from data sheets. For older Ig's, can drop the dilution (e.g. from 1/1000 for anti-b-actin, to 1/500.
2. Prepare dilution solution. We couldn't find any goat serum so used BSA (bovine specific albumin) instead. The protein just prevents non-specific binding later on. We dissolved 100mg BSA in 10mL PBS (1x) at room temperature.
3. Using a disposable pipette, draw off excess PBS from relevant wells in 24 well plate. Add in 200 microlitres of PBS/BSA solution to each of the relevant wells. Leave for a few minutes whilst completing next few steps.
4. Set up a piece of parafilm on a glass plate - stretch slightly so it sticks well. On the parafilm, use a marker to draw in a grid consisting of the cells being used, whether they are differentiated or undifferentiated (and a column for the control) - there should be a space for each coverslip being used such that the slips can easily be identified once taken out of the 24 well plate.
5. Make up primary Ig solution in an eppendorf tube (remember, that we are double staining, so both Ig's go in the same solution and at the same volume). i.e. To 500 microlitres of PBS/BSA solution, 1 microlitre of anti-B-actin MAb was added (1/500 dilution) and 5 microlitres of anti-tau PAb was added (1/100 dilution).
6. Mix Ig solution with 200 microlitre pipette - no bubbles.
7. Add a 30 microlitre drop of the Ig solution to the relevant grid space in the parafilm grid.
8. Remove the relevant coverslip from the 24 well plate using a needle and forceps (using needle to get leverage) and be sure to remember which side the cells are on. Place the coverslip cell side down onto the drop of Ig solution on the parafilm.
9. Repeat steps 7-8 for all relevant coverslips. HOWEVER, for control, place the slip onto a drop of PBS/BSA (NOT Ig solution).
10. Cover up the slips with a plastic lunchbox (they must not dry out) and let them incubate for 1hr-1hr 30 minutes (depending on the room temperature).
11. Prepare secondary Ig solution (again, both secondary Ig's in the same PBS/BSA volume) to appropriate dilutions. Ensure you have enough for 30 microlitres per coverslip.
12. Prepare a new grid of parafilm on a new glass plate.
13. Ready three 100mL beakers of PBS (1x) for washing of coverslips. Also have some wads of kim wipes ready to dab off excess solution/PBS.
14. CONTROL COVERSLIPS ARE WASHED FIRST! Have a drop of secondary Ig solution ready on the parafilm grid. Remember cells are face down when you pick up the coverslip with forceps. Dab of excess solution by touching side (ONLY) of coverslip against kim wipe. Rotate coverslip if necessary. Dunk coverslip in first PBS beaker and dab off excess again. Repeat 5 times before moving to next beaker. Repeat 5 times also in second and third beakers. On last wash in the third beaker, be careful to remove as much PBS (by dabbing on kim wipe) as possible - as otherwise it will further dilute the Ig.
15. Drop the coverlip cell side down onto the drop of secondary Ig solution on the parafilm. Repeat the washing process for all coverslips.
16. Leave incubating in a cool room temperature overnight, INSIDE a plastic lunch box lined with moistened (RO water) paper. i.e. put the whole glass plate and coverslips in the lunchbox and put lid on.
17. Next morning, rinse coverslips again with the three beakers of PBS. Dab off excess and this time place on drops of PBS/BSA on a new parafilm grid.
18. Prepare slides by wiping clean with kimwipe and labelling with name, date, cell line, Ig's and colours.
19. Make up the 'ProLong solution'(Prolong Antifade). This can either be done by the provided protocol if all the solution will be used up at the one sitting. If however, the normal protocol produces too much solution for your purposes (10-15 microlitres are needed per coverslip), you can instead use a solution A that is mixed in 1005 ethanol. Then mix this with solution B. The ratio is A:B, 1:10. Eg. for 11 coverslips, need 165 microlitres. Therefore, use approx. 160microlitres of B added to 16microlitres of A. NOTE: The solution should only be made immediately prior to using it - it shouldn't be stored.
20. Place 15 microlitre drops of solutions onto slides, pick up coverslips and dab off excess PBS/BSA and drop onto solution. Leave enough space on each slide for up to 3 coverslips.
21. Leave slides covered with plastic lunch box, to dry and set (the ProLong solution turns to a gel-like consistency, but the coverslips must not be moveable at all). This can take between a few hours and overnight.
22. Wash off PBS crystals once the antifade gel has set firmly. Do this by dabbing a wet (RO water) kimwipe over the slip.

Notes on using the confocal microscope

These are only notes, not full instructions. An induction must be carried out on this Olympus FV1000 with Gavin McKenzie at the HMU unit, UNSW. There is also a folder next to the microscope which details some of the following notes.

1. If you are the first user of the day, turn on the microscope components from left to right. That is, turn on the lasers (black boxes) first by flicking the key AND switch on each., skip the UV light box if not required, turn on the power box for the mercury lamp (3rd box), skip the next and turn on the power box at the back. (The latter 2 are large, white boxes). Next switch on the white box next to the CPU and turn on the CPU itself.
2. Log into the CPU (password not given here) and open the FV10-ASW program.
3. Wipe down the stage/objective with a kimwipe and 70% EtOH.

Notes on the 'Acquisition Setting' Window - Scan speed - faster = less contrast but also less bleaching. Vice versa. Good trade off is 10us/pixel. Optimisation of images is done at the fastest speed of 2us/pixel. - Image size - 1024x1024 is best - Area - magnification tool is useful to zoom in at a certain magnification without changing objectives. Just click the magnifying glass, highlight an area in the live window (whilst it IS NOT scanning), then scan the image. It will only scan the zoomed in area. - Lasers area auto adjusted (no need to change). 405 at 20%; 488 at 25%; 543 at 45%; 633 at 45%.

Notes on the 'Image Acquisition Control' Window - Green box allows to choose dyes as required for the slide/s. (Eg AF488 and TRITC) - Turn on the mercury light/lamp using the 'EPI' button.

Notes on the Control Panel to the right of the microscope - Filter cubes indicated by WIG (use it to see red dyes such as TRITC and Rhodamine); WBV and NIBA (use to see green dyes such as AF488).

1. Depending on the power you wish to use, the objective will require a drop of water or oil to be placed on prior to putting the slide on (refer to the name of the objective in the CPU program. The last letter will indicate water or oil (o).). NOTE - SLIDES MUST BE PLACED UPSIDE DOWN ONTO THE OBJECTIVE!
2. Once the slide is placed on and held in place by the holders, move the stage up (using the electronic buttons/arrows on the control panel) until the slide just gets slightly pushed upwards.
3. Turn on the mercury light on using the EPI button as stated above.
4. Should see a haze of fluorescence. Tap the down arrow focus button until you see cell outlines. Then fine focus to see the cells clearly.
5. Find the cells you are interested in and then turn off the mercury lamp using the same EPI button.
6. NB When using >1 fluorescent probe, must click 'sequential' function button to account for bleed through. Make sure sequential is checked.
7. Click '2x scan' button in image acquisition control window (at top). Whilst it scans, fine focus the cells in each dye. This is the first step of optimisation. Note that 2x scan is a faster scan that only gives a rough image. More detailed images will be given during image capture
8. Optimisation step 2 - move the voltage (HV bar in the image acquisition control window) up and down to get the best signal (not too bright). NB: Press Ctrl + H on the keyboard to get saturation image - this will help to perfect the brightness (indicated by red dots) and the offset (which should be altered until there is a clear focal contrast between the black background and blue cells. Regarding the brightness, you don't want too many red dots).
9. To capture the image, use the 'XY' button, NOT the 2x scan.
10. The image will open in a new window. To save it, right click >> Export >> save as Tiff. The third button from the left (at the top) will show an overlay for when you have captured >1 dye. Use the pencil tool to add a scale bar. NB: If you add a scale bar, DO NOT use export function to save. Instead, use 'Save Display'.
11. NB: If you use the 'save as' function in the right click menu, it will save as an OIB file that can only be opened with this program. It is worth creating an OIB for your negative controls (to which you compare all your other slides) because saving as an OIB also saves the optimisation data (brightness, offset, gain etc). To make these settings the same, when you open the OIB file, click the '<<' button at the top left of the 2D image window. Click the tools button on the far right and let the settings re-set.
12. You can use the TDI to get phase contrast images - Adjust the brightness. (This function does not work that well with out cells?)

Following use - if you are the last person of the day to use it, turn off everything in reverse. For the last black box that has a SWITCH and a KEY, leave the SWITCH ON (it is a fan), but turn the KEY OFF

Realigning the Leica DMIL Microscope Mercury Lamp (106z lamp housing)

See pages 52-54 of the manual also.

1. Switch on the mercury lamp and open the light stop on the left.
2. Move a filter block into the light path.
3. Coarsely focus the light with a low to medium magnification, on a sheet of blank, white paper placed on the specimen stage.
4. Mark the centre of the bright surface on the paper, with a pen.
5. Remove the objective.
6. Rotate the collector adjustment (front knob on left of lamp housing)until the lamp filament is clearly projected.
7. Move the reflection of the lamp filament to the side by rotating the top and bottom knobs at the back of the lamp housing (NOT the middle knob).
8. Focus the direct image of the lamp filament by first using the horizontal and vertical adjustment controls of the holder (the back knob on the left of the lamp housing and the knob on top of the lamp housing)to move the direct image into the centre of the brighter circular area. Then move the reflection into the brighter circular area. Focus the reflection and asjust the mirror until the reflection is super-imposed to the direct image.
9. Remove the paper and test using a slide. The light illumination should be homogenous.