protocols in biomedical neuroscience

PROTOCOLS IN BIOMEDICAL

NEUROSCIENCEPrepared by Kasra Houshmand

Temple University Medical Education Research Building7th floor – Neuroscience, zone 3

Kasra Houshmand, Biomedical Neuroscience Protocols 1

Recipes 10% FBS – DMEM

500ml bottle of fresh DMEM media (cell culture fridge)50ml of FBS (fetal bovine serum) (cell culture fridge)20 ul Gentomycin

1.5 M Tris, pH 8.8Make a 1.5 molar solution of Tris HCL (mw = 157.6), then pH the solution to 8.8 using a basic buffer such as 5.0M NaOH.

1.0 M Tris, pH 6.8Make a 1.0 molar solution of Tris HCL (mw = 157.6), then pH the solution to 6.8 using a basic buffer such as 5.0M NaOH.

10% SDS5.0g Sodium Dodecyl Sulfate + 50ml dH2O

10% APS1.0ml dH2O + 0.10g APS (chemical room – APS should be in 4 degree and does not last many days)

Running Buffer: 10x TGS Running Buffer:

First, fill a 1000-2000ml graduated cylinder up to 500ml dH2O+30.2 g Tris Base (chemical room in the floor level cabinet)+144g Glycine (chemical room on the floor under the bench)+10g SDS…allow everything to dissolve-Fill to 1 liter dH2O

Transfer Buffer:1x Transfer buffer (do not make 10x stock transfer buffer)

First, fill a 1000-2000ml graduated cylinder up to 200ml dH2O+3.1g Tris Base (chemical room in the floor level cabinet)+14.4g Glycine (chemical room on the floor under the bench)-Fill up to 800ml with dH2O -Fill up to 1liter with Methanol (200ml methanol)

1x PBST100ml 10x PBSTUp to 1000ml with dH2O (1:10 dilution)

5% Non Fat Milk o Blocking solution / buffer for antibodies

30ml 1xPBST 1.5g Non – fat dry milk (or relative proportion)

Primary antibody mix in 5% NFMo 1:1000 ratio usually, so 15ml of 5%NFM, and 15 µl primary antibody

Secondary antibody mix in 5% NFMo 1:5000 ratio usually, so 15ml of 5%NFM, and 3 µl secondary antibody (Agno protein gets RHRP,

a polyclonal antibody.)


Transfection Bring out DMEM / Optimem to bring to room temperature 60µl Optimem + 6 µl Lipofectin + 2 µl DNA (example: CMV-Agno) in an eppendorf tube. DNA

plasmids have set concentrations; you must calculate how much DNA there is per µl to put in 2 µl total. (example: CMV-Agno is 0.5 µl DNA / µl – so you need 4 µl total to get 2 µl DNA from this sample)

Primary incubation of 45mins (inside the hood is fine – just leave the eppendorf tubes closed on a rack)

Add +500 µl Optimem to each eppendorf tube after incubation Transfer the mixture dropwise to the plated cells, after aspirating the medium. If there are any

untransfected cells, leave them in their normal medium for now, at the end of the process all the cells, transfected or not, will receive new medium.

4 Hour incubation in the incubator – softly shake the plates every 30 minutes. Aspirate transfection solution at the end of incubation Replenish with fresh medium If there are untransfected cells also involved in the experiment, replace their -medium at this time

as well by aspirating / replacing

Whole Cell Extract (for 6 plate dish – amounts of Trypsin / media is different depending on plate size)

Aspirate the old medium Trypsinize (+0.5 µl TrypLE), incubate 2-5mins, check to make sure cells are released Add fresh medium (+2.0 µl fresh DMEM) to neutralize trypsin. Centrifuge cells @ 1200 RPM, 5 minutes. Aspirate supernatant. Resuspend pellet in 1 ml PBS, transfer to an eppendorf tube, and then microfuge cells @ 7000

RPM, 2 minutes. Aspirate PBS Resuspend in TNN buffer (1% TNN with MPI…mammalian protease inhibitor) MPI: TNN ratio is apx. 1:100, (10 µl MPI in 1000 µl TNN) Each sample gets apx. 400 µl TNN (DEPENDS ON PROTIEN CONCENTRAION – sometimes

400 µl is too much) Rotate @ 4 degrees for 45 minutes Spin @ max (4 degrees) for 10 minutes Save supernatant in new tube; discard pellet LABEL PROPERLY

Extracting Cell Medium for extracellular studiesKasra Houshmand, Biomedical Neuroscience Protocols 3

Pipette the old medium into a 50ml tube. (do NOT aspirate, it’s being saved) Spin at 3000RPM, 5 minutes. Aliquot the supernatant into a new 50ml tube, freeze at -20 degrees C. Trash the pellet.

CO-IPDay 1: 4ml medium + 25ul beads + 5ul NRS / Antibody + 6ml TNN buffer Rotate at 4 degrees C overnightDay 2: Centrifuge @ 3k, 5 mins Aspirate the sup / keep the beads (pellet) Resuspend the beads in 1ml TNN then transfer to an eppendorf tube Centrifuge 7k, 1 min Aspirate the sup / resuspend in 1ml TNN buffer, then centrifuge 7k, 1 min. Do once more…aspirate the sup / resuspend in 1ml TNN buffer, and then centrifuge @ 7k, 1 min.

(a total of 3 TNN resuspension / centrifugations) Last wash – replace TNN with PBS – then aspirate the PBS after 7k, 1 min centrifuge. Resuspend beads in 50ul dH2O after aspirating PBS Add 10ul 5x SDS dye Boil @ 95 degrees C for 10 min Run on WB gel

ICC: Immunocytochemistry


Fixing steps: Plate cells on chamber slides Each chamber has its own condition – for example four plates can be fixed at increasing time

points, such as 9, 12, 24, 48 hours. Fixing solution: ice cold methanol / acetone solution is used to fix the cells, essentially freezing

all activity and allowing for visual analysis up to months afterward. First, aspirate the condition media of the cells Drop in ~0.5 ml fixing solution to the chamber (the chamber has a maximum capacity of about

1ml) Let it sit for 30-45 seconds, and then add PBS to the fill line. (NOT PBST used for WB

membranes, but PBS, used in whole cell extract) Aspirate the entire chamber, making sure to keep the aspirator tip near a corner so the cells

remain undisturbed. Wash with PBS by filling the chamber, and then aspirate. Fill chamber with 0.5ml PBS, and then allow it to sit. At the end of your experiment, however

many chambers there are, you will replace all the PBS with fresh PBS once the last slide has been fixed and washed.

Washing steps: much like western blot membranes Lay the chamber slides in a box with PBS soaked napkins underneath so the slides don’t shift

around. Fill each slide up with PBS, wash 1x, and then dump contents. Blocking: 5% BSA (bovine serum albumin) in PBS, 1 ml per chamber slide. Block for 2 hours on

rocker, in box. Dump contents – wash 3x for 15 minutes each with PBS Primary antibody: 2.5% BSA in PBS, with 1:500 antibody – run overnight Dump contents – wash 3x for 15 minutes each with PBS Secondary antibody: for 8 slides, 8.8ml PBS with 10 µl secondary antibody. Put 1ml of the

master mix into each slide, covered in the DARK. (a covered box will do) Probe secondary for 2 hours.

Dump contents – wash 3x for 15 minutes each with PBS

Mounting steps: Remove the top part of the chamber slides with the removal tool provided. The slides, where the

cells sat, is the only required item now. Apply mounting media to each slide – about 2-3drops of DAPI mounting media per slide. Lay down a clean glass on top of the slides – place down carefully so there are no bubbles. Once covered, tip the slide lengthwise on the bench, allowing the excess mounting media to drip

onto a Kim-wipe underneath. Glue the glass down to the slides, as shown in the image. Nail polish is used as the adhesive, glue

down all four sides of the glass to the slide. Remember – once secondary antibody is applied, you need to keep the slides in the dark. Do not

expose the slides to light for too long after secondary, work quickly when in light. The slides are now ready to be examined under the microscope.


ICC: Aspirating first chamber to fix cells

ICC: Slide portion of chamber slides, (chambers have been removed) covered with glass and glued down after applying DAPI mounting media. These slides are now ready for analysis.


Bradford Protein Concentration Assay

Write down the equation from the bottle for concentration calculation later on +1ml of Bradford dye per cuvette +5 µl of protein sample per cuvette Note: proteins turn the dye blue from its original tan color. More protein means more blue

color. Read each sample in the 595nm spec, auto zeroing a “blank” cuvette with only 1ml of dye in

it Take the absorbance values, also known as the “OD” value, and plug them into the equation

provided on the Bradford dye bottle to get [e], the protein concentration. This concentration is what determines how much sample is needed for each well in Western

Blot Western Blot calculation: we usually want 60 µl total sample size per well (this is including

water, dye, and protein sample) The main factor is how much protein sample is needed. You divide the desired amount of

protein, in micrograms, by the concentration…for example:

If we want a 50 µg sample of protein 1, and our concentration of the protein is [e] = 2.0 µg/ µl, then we would do the following calculation:50 µg / 2.0 µg/ µl = 25.0 µl protein sample

Since we usually want 60 µl total sample size, the remaining amounts would be as follows. 10 µl SDS running dye is always consistently added to the samples, regardless of desired concentration.

Water Dye SampleLadder - - 10 µl (no

water/dye for ladder)

Protein 1 25 µl 10 µl 25 µlProtein 2 10 µl 10 µl 40 µl

This chart suggests that protein 2 has a much lower protein concentration than protein 1, as you need 40 µl of protein 2 to get the same amount, in µg, of protein given by sample 1 at 25 µl.

Western Blot


1. Making a gelLarge glassSmall glassGlass clamp2 Spacers (large thickness, make sure they are same thickness)Well combLeak-proof (ideally) gel-making mount (1)

Fit the BioRad WB aperture together: Large Glass in back, spacers, and then small glass. Check for leaks using dH2O. (1, 2, 3)

Make resolving solution while checking for leaks – recipes vary, check with your PI. Pour 7.5ml of resolving gel into the space between the glasses, and then pour dH2O on top.

Once the resolving layer has polymerized, a line will appear. It usually takes 15-30 minutes to polymerize fully. Check the remainder of the solution in your plastic tube to see how it is progressing. (4)

Remove the layer of water on top by simply tipping the aperture over, and then remove the remaining water with chromatography paper, it will absorb only the free water. Ensure there is a solid straight line of polymerized gel – sometimes leaks cause the sides to fall down below the straight line slightly…if this is the case, avoid loading samples into either end of the gel, as they will run mildly sideways and not look good. (4)

Make the stacking gel and pour it on top to the top. Immediately place the comb inside, slightly overlapping the front of the two black spacers. As a point of reference – you must leave stacking space underneath where the wells end in the stacking layer, such that there is a straight layer of stacking still in between the wells and the resolving layer. The space between the wells of a 10-well comb is about 3-4 millimeters, this is approximately the space needed for the stacking layer – so ensure when placing the comb, that you do not leave too thin of a stacking layer, or else the proteins will not accumulate well to run down the resolving layer. Wait for the stacking layer to polymerize. It usually takes 15-30 minutes to polymerize fully. (5)

Once the stacking layer has polymerized, remove the comb by slowly and steadily pushing the comb up and out of the wells. (5)

Clean the gel wells off with running distilled water to ensure no pieces of polymer are stuck inside the well. To fully extract anything left inside the wells, calmly – but firmly – whip the gel aperture away from you to send water and excess gel particles flying out of it…use a whipping, steady wrist action to clean the wells out.


1: Bottom – small glass, large glass; middle – white well comb, two black spacers; top – gel making mount, glass clamp. (read left to right)


2: left: a fully installed glass + clamp set. Right: a “blank” clamp, used when only one gel is being run at a time, since there are two slots in the electrophoresis box.)

3: clamp with fully installed glass set-up, mounted on the leak-proof gel making mount)Kasra Houshmand, Biomedical Neuroscience Protocols 10

4: polymerized resolving layer

5: comb placed in stacking layer to polyermize


2. Sample loading

Electrophoresis box / wiringPower Supply1x SDS Running buffer

Place the polymerized gel, still mounted inside its clamp, on a electrophoresis mount that will be placed inside an electrophoresis box. (6,7) If you have only one gel to run, you will load a “blank” clamp, tightened down all the way, on the other side. Also, pay note to image 7 – the rubber stopper in the aperture can be flipped and used on both sides – the flattened side is for blanks to ensure there are no leaks. The other side is for loaded gels. The fit is different for the two, so be sure to have the rubber facing on the proper side. Bumpy, darker side: actual gel. Flat, lighter grey side: blank clamp.

Prepare your samples in eppendorf tubes – always make a chart like shown on page 1 to display how much sample, water, and dye are being used in each well. Label your wells in your notebook consistently.

Boil your samples for 5-8 minutes, and micro-centrifuge the samples quickly at max, then the samples are ready to be loaded. Boiling and the Beta-mercapto ethanol in the SDS running dye are done to denature the proteins.

Pour 1x SDS running buffer inside the box to level, ensure there are no leaks, and load the wells, using a protein loading tip on a P200 micropipette to load the entire sample at once.

Run at 150volts through stacking phase, and turn voltage up/down at resolving phase to specifications. (8, 9)


6: blank glass clamp attacked to the box mount with the prepared, filled clamp ready to load to the other side

7: this grey rubber stopper on the electrophoresis mount has two sides that can be flipped – this side, the lighter grey, flat side, is used to attack blanks to, ensuring no leaks in the running process. You must flip this rubber piece over for mounting a loaded clamp.


8: electrophoresis set-up, with box, wiring, and power source

9: samples loaded, gel running


3. Transfer phase

Recipes / reagents:1x SDS Transfer BufferMembrane (either PVDF or nitrocellulose)Gel from WBTransfer Aperture – black / clear cassette, transfer box, wiring, and power source

Clear SideSpongePaper

Gel – facing upMembrane

PaperSponge

Black Side

(Have the black side of the cassette face the red side of the transfer box) (10, 11) First, wash the membrane (with methanol. If a PVDF membrane, with dH2O if a

nitrocellulose membrane). Immediately wash off the methanol with dH2O until the water does not bead on the membrane and completely runs off, leaving no residue behind.

Load the cassette in the mentioned stacked order, submerged in transfer buffer in a large glass pan. Once done stacking, ensure there are no bubbles in the sandwich by softly pressing out from the middle to each side, but carefully holding the stack in place so that nothing moves around inside – if the membrane does not fully cover the gel, you will not get a full image of the blot.

Load the cassette into the transfer electrophoresis mount, place it inside the electrophoresis box (same box used for running), fill with 1X transfer buffer and place a stir bar at the bottom of the box. Place the entire box on a large glass pan to ensure no spilling.

Run in the 4 degree cold room, time and voltage depends on protein. (Agno takes 10minutes, a relatively short run for transfer) Run the stir bar at a low speed, the voltage is usually 250milliamps.


10: stacking sandwich setup. The clear side is hanging off the edge over the left side, with a sponge sitting on top. The right side of the stack is submerged in transfer buffer, in the following order from bottom to top: black side, sponge, paper, membrane, gel (facing UP, with the ladder on the left side, since it was the first well loaded in line). On top of the gel will go another paper and the sponge as shown, and then the sandwich will be closed, after ensuring, under submersion, that there are no are bubbles remaining – to remove bubbles from the sandwich, hold it carefully in place and slide the flat back of your finger from the middle towards the edges to send bubbles out. At this point, the sandwich is ready to be placed inside a transfer electrophoresis aperture with the black side of the cassette facing the red side of the transfer electrophoresis mount.

NOTE: the electrical aperture runs from black to red. This means that the cassette, which is oriented with its clear side closest to the black side of the electrical aperture, will run from clear to black, just as the aperture runs from black to red. This means that the proteins which are on the gel will run towards the membrane, as according to the order of the sandwich stacking.


11: This image shows a cassette, loaded into a transfer electrophoresis mount, inside an electrophoresis box. (the same box used for running is used for transfer, simply a new aperture is placed in it) Note that the black side of the cassette is facing the red side of the mount, which means that when the mount is running from black to red or also known as negative to positive, the materials inside the cassette sandwich are moving from clear side to black side, with the gel closest to the clear side, and the membrane closest to the black side of the cassette, therefore transferring from gel to membrane.


4. Probing antibodies

Recipes / reagents: Blocking solution: 5% non fat milk in 1x PBSPrimary antibody: 1:1000-3000 (antibody: 5% NFM) Secondary antibody: 1:3000-1:5000 (antibody: 5% NFM) Block for 40 minutes with blocking solution Note: everything is done on the rocker for blocking, washing, and probing. Wash 1x with PBST, 5 mins Probe primary antibody for 3-4 hours Wash 3x with PBST, 5 mins each Probe secondary antibody for 45-1 hour Wash 3x with PBST, 5 mins each Once the membrane is at this point, it’s ready to prepare to be developed in the dark room.

5. Developing in the dark room

Recipes / reagents:Developing reagent (commonly ECL or ECL+, need to follow the specific reagent instructions for ratios of reagent A + reagent B, and the total amount needed per membrane area in centimeters squared)X-ray cartridgeX-ray film

Lay down saran wrap on the bench totally flat using water to assist in flattening. Use a Kim wipe in an outward wiping direction to flatten.

Lay down the membrane totally flat, and apply the developing reagent. Lay a piece of parafilm over top of the membrane to ensure full coverage.

Fold the saran wrap over top of the membrane after short incubation and once again ensure there is a flat seal over both sides of the membrane by wiping any bubbles or folds out of the plastic. Cut the plastic to size, and tape the laminated membrane inside a film cartridge – it’s best to mount it to a corner so you can use one piece of film twice by simply flipping it to expose the other half.

Take the cartridge / film to the dark room and develop. Load film in total dark; usually use one film for two exposures.

It is best to develop a system such as justifying the membrane to the top left of the cartridge, and then justifying the film to the same corner, and then simply flipping it to expose the other half. It also helps to fold a corner of the film to remember what the orientation is for yourself.

Usually the first two exposures are 30 seconds, and 1 minute, depending on the anticipated signal that will come from the membrane.

Run the films through the developing machine; turn it on if it’s not already running. Once you’ve seen how well the image was exposed, you may estimate how much longer/shorter the next one needs to be – but work fast, the signal will fade.


Direct ELISA

Direct ELISA Protocol – agno specific(modified, source: abcam)Kasra HoushmandIlker Sariyer11/18/11Temple University, MERB, Neuroscience, 7th floor, zone 3

Solutions: 1. Antigen coating buffer2. α-Agno antibody (7903 IsG)3. Blocking Buffer: 2.5% BSA, 2.5% milk4. PBS (phosphate buffered saline)5. Antibody dilution buffer: 1x blocking buffer – agno antibody dilution 1:5006. Substrate solution (TM blue)7. Stop solution

Steps:1. Antigen coating: 50 µl sample/standard + 150 µl antigen coating buffer. Mix, incubate overnight

at 4 degrees. 2. Wash wells with PBS = 200 µl each, 2x3. Block the wells with 200ul blocking buffer = 2-4hours at room temp 4. Wash 2x with PBS5. Add 150 µl of primary antibody into the wells6. Incubate overnight at 4 degree shaker7. Wash 3x with PBS (5min washes)8. Add 150 µl of secondary antibody (α R-HRP, 1:1000 dilution in 1x blocking buffer)9. Incubate 2-4 hours at room temp 10. Wash the plate with PBS (200ul/well) 4x, 5 min washes11. Add 100 µl of substrate solution into the wells – let color develop12. Add 100 µl of stop solution13. Read OD’s in plate reader (455nm wavelength)14. Analyze data


MBP Bacterial Work

MBP Protein WorkKasra Houshmand09 November 2011

Day 1: Growing culture/collecting pellet1. Grow 1L culture with LB + AMP (1L LB : 1ml AMP ratio)

Some times you will start with a 200ml culture, remove 100ml of it and then add 900ml fresh LB+Amp media to produce more protein. This 100ml that you remove can be used to make glycerol stocks for freezing. (Glycerol freezing media: 50% glycerol, 50% LB+Amp. When freezing culture, add 500ul culture, 500ul freezing media.)

2. Shake culture at 37 degrees, check occasionally for growth.3. Read OD’s @ 595 nm – once the levels reach 0.6-0.7, change to 28 degree shaker. After 30

minutes, add IPTG to induce protein production, (500ul IPTG to 1L culture) then continue incubating in 28 degree shaker for two more hours.

4. Centrifuge in large high speed tubes @ 8k for 10 minutes, 4 degrees5. Resuspend pellet in 20ml COLUMN BUFFER

(COLUMN BUFFER: 200mM NaCl, 20mM Tris HCl, 1mM EDTA. Keep at 4 degrees…make 500ml at a time)From here on out, always keep protein samples cold, and column buffer is basically used for every step here on out.

6. Sonicate the suspension – apx. 3 10 second sonications should be adequate. Avoid frothing – lysis is complete when cloudy suspension becomes translucent.

7. Centrifuge in small high speed tubes @10k for 10 minutes, 4 degrees8. Transfer supernatant to 50ml capped tubes, trash the pellet.9. Add 100-300ul Amylose resin beads, put on 4 degree shaker over night.

Day 2: Running on SDS Gel / elution1. Centrifuge tube @ 3k for 6 minutes2. Resuspend pellet in 1ml COLUMN BUFFER – transfer to eppendorf tube3. Wash beads 4-5x with column buffer, resuspending each time gently and then centrifuging. 4. Resuspend final time in 150-200ul COLUMN BUFFER5. Take Bradford concentrations6. Run on 10% gel, allow all dye to exit before stopping.7. Stain gel with coomasie blue, destain.

Purification Steps:

ELUTION BUFFER: 10mM maltose in COLUMN BUFFER (make 50mls at a time)

1. Spin 10k 1 minute, remove supernatant. (eppendorf tubes)2. Wash beads with COLUMN BUFFER one last time, spin @ 10k for 1 minute.3. Remove supernatant. Add 60ul of ELUTION BUFFER. Allow sample to sit at room temp for 5

minutes. Gently resuspend, then centrifuge @ 10k for 1 minute. Take 50ul of the supernatant (leaving 10ul behind) and transfer to a labeled tube “elution 1.” Add 50ul of ELUTION BUFFER back in to the tube, and repeat the 5 minute room temperature incubation /


centrifugation. Once again transfer 50ul to a labeled tube, “elution 2,” leaving 10ul behind. Do this step 4-5 times, each time saving 50ul of the elution.

4. Read the OD’s of these elutions in Bradford assay – combine together the tubes that have the highest protein concentrations, and once again read the OD of the master mix. Toss the elutions that have no proteins. For the blank here, use 1ml Bradford dye + 5ul ELUTION BUFFER. For the samples, use 1ml Bradford dye + 5ul elution (suspended in ELUTION BUFFER)

5. This purified protein is now ready to be run on another 10% SDS gel, as purified proteins.

Source:“Small and Large Scale MBP-fusion Protein Purification”The Protein Purification Facility The Wolfson Centre for Applied Structural BiologyThe Hebrew University of Jerusalem Dr. Mario Lebendikerhttp://wolfson.huji.ac.il/purification/TagProteinPurif/MBP_Tag_nature.html


http://wolfson.huji.ac.il/purification/TagProteinPurif/MBP_Tag_nature.html

protocols in biomedical neuroscience

Documents