Autoradiography 35S

  • Remove Kodak NTB2 nuclear emulsion from fridge and place at 42oC for around 30-60 mins (until melted).
  • Make up the developer and the fixer and place in the darkroom ready for later (only if developing the same day). The developer should be a 1:1 dilution in water. The fixer is 30% (w/v) of sodium thisulphate.
  • Place slides in a dark box and carry to dark room. Also take a slide rack, tweezers and the emulsion.
  • Switch on the red light and switch off the main light.
  • Working with the red light, unwrap the foil from the emulsion. Place the emulsion in the slide rack.
  • As quick as possible dip a slide into the emulsion with the tweezers. Place the slide in the slide rack to dry. The quicker the slide is dipped, the thinner the coating of emulsion and the better the visualisation later.
  • Switch off the red light.
  • After around 30-60 minutes the slides will be dry. If you don???t want to wait in the dark for half an hour make sure to place a note on the darkroom so people know not to enter.
  • After the slides are dry, re-enter the room but??don???t turn on the light. Working in the dark, place the slides into the dark box and wrap in foil. Also wrap up the emulsion.
  • Store slides in a fridge or cold room

??Developing the slides:

  • Remove slides from fridge and allow to warm to room temperature for around an hour
  • Working in the dark, place the slides in a slide holder and immerse the slide holder in D19 developer from 3 minutes. Time with a countdown timer as you won???t be able to tell the time.
  • Immediately place the slide holder in the fixer. Leave for 3 minutes again.
  • Turn on the main light.
  • Wash the slides with running tap water for around 5 minutes.

Formaldehyde Treatment Of Tissue Culture Hoods

You will need:-

12g Potassium Permanganate??
6g Crushed paraformaldehyde

1) Set the hood so that it can vent outside.

2) Mix the two chemicals above together (in a jam/coffee jar, so it can be thrown away) in the hood, wear gloves and a face mask.

N.B.:??Potassium permanganate is a??strong??oxidising agent while paraformaldehyde and formaldehyde vapour are??extremely??toxic.

3) Add 50ml water and mix.

4) Quickly place the front door of the hood in position, and tape over the joints.

5) Leave overnight and then switch on hood for at least 1 hour prior to use.

A Protocol For Cleaning And Reusing The Large 25 X 25 Cm Plates

We regularly reuse our large 25×25 cm plating trays; initially, however, we were plagued by gross microbiological contamination when reusing the trays. Various combinations of cleaning with a laboratory dishwasher, ethyl alcohol, and UV irradiation still resulted in a lot of contamination, particularly yeast. After examining several disinfection methods, the following procedure was determined to be the simplest and most effective way to control microbiological contamination of previously used plating trays.

1. Scrape out the old used agar/agarose into autoclave bags. Autoclave and discard. Adding a spoonful of baking powder prior to autoclaving is reputed to significantly reduce the sometimes foul smell that occurs before and after autoclaving [recommended by D. Stout in a letter to ASM News, 59(#2) : 51 (American Society of Microbiology, 1993)]. Try using approximately two tablespoons per 10 liters.

2. Wash the empty, dirty plates under the tap, bottoms and tops, using a soft sponge (no scouring pads or abrasives!) to ensure debris is removed from the plates. Be careful to ensure that any yeast or other visible unwanted material is removed.

I also keep a large plastic beaker nearby with 95% EtOH and a wad of paper towels or a cloth, for removing marker pen markings from the plates as I am washing them at the sink. After washing the plate, wash off any ink with the EtOH, then wash away the ink/EtOH with a final rinse under the tap.

3. Stack the plates neatly at the side of the sink as you finish washing them.

Note : Always ensure that plates are neatly vertically stacked and aligned. The weight of a stack of poured plates (200-250 grams/plate) is considerable, and any misalignment when they are stacked will eventually lead to cracked lids, due to pressure points bearing the full stack weight load. For the same reason, I try to limit stacks to 20 plates or less. Of course, another potential disadvantage of improperly stacked plates is warpage (particularly if the plates are heated – see item #8, below).

4. Once you have a suitable number of tap water-washed plates, transfer them to a pre-cleaned (scrubbed to remove obvious debris) sink, filled with room-temperature water containing approx. 1% bleach (Clorox; hypochlorite). For our sink, I fill the sink about 3/4 full (approximately 50 litters) and add 500 to 1000 ml Clorox. (500 ml gives a final concentration of about 1% bleach). I let the plates soak in the dilute bleach solution for approximately 1/2 hour or so (while I am scrubbing another set of plates under the tap). Our sink holds roughly 40 plates at a time, standing vertically, with the lids in place over the bottoms. I am careful to ensure that the plates are completely submerged, and that there are no bubbles shielding any of the plate surfaces from the bleach.

Notes : (a) I strongly recommend the use of a lab coat, gloves and eye protection while working with concentrated and dilute bleach solutions. (b) Halogens will slowly eat away at stainless steel, as anyone with HPLC experience should already know (columns are flushed free of halogens at the end of the day). Therefore, I would recommend rinsing the sink with tap water, after use with bleach solutions.

5. After the plates have soaked, grab a small stack of about 4 or 5 plates (with the lids in place over the bottoms), and let the bleach drain into the sink. Gently shake excess bleach from the plates, holding the stack of 4-5 plates together with the lids in place. Place them on a trolley. Fill the sink with the next set of plates to be bleached.

6. Rinse the bleached (disinfected) plates with Millipore NANOpure 0.45um-filtered ddH2O (I use the water from our ddH2O storage/wash tank next to the sink). Rinse the bottom plate first under flowing ddH2O (both sides), then rinse the corresponding lid and place it over the bottom plate. The ddH2O rinse (of course) is to wash away residual debris and sodium hypochlorite.

7. Again, once you have a stack of 4-5 bottoms with the lids in place, grab them and shake out the excess ddH2O. Keeping the lids on the plates will help minimize airborne contamination.

8. As a test, I let several of these plates (4) sit overnight, then poured bottom agar. They all seemed fine. However, for most of my plates, I further incubate the plates at 70C overnight (in stacks of 20, unbagged), taking care to ensure that the plates are perfectly aligned vertically (no pressure points). The 70C bake may not be necessary in terms of disinfecting, but it is a quick way to dry the plates. The plates appear to survive baking at 70C with no adverse effects.

9. Stack the plates until ready for use (pouring bottom agar) on the bench; I cover mine with plastic, to minimize airborne contamination (I have not examined whether this helps or not).

10. Pouring the bottom agar – as per usual methods. After the bottom agar has solidified, I invert the plates (bottoms up), set the bottom aside, and wipe excess condensation from the lids using ‘giant’ lint-free KimWipes. This helps prevent water droplets (rivulets; rivers) on the agar surface and at the edges of the plates, minimizing the spread of random contaminants on the agar surface (some low level random contamination will occur if you pour plates on the bench; while this will be minimized if a laminar flow hood is used, in our experience it is unnecessary). Incubate the plates at 37C overnight. Carefully check the for microbiological contaminant growth (there should be very little). Bag the plates (into clean bags) and keep them on the bench (up to several days), or refrigerate until use.

Notes : (a)The procedure of wiping the lids with KimWipes doesn’t appear to introduce any significant degree of contamination. (b) I would recommend using the plates within a few days of pouring, the sooner the better. (c) After an overnight incubation at 37C, there may be a few colonies on some of the plates. This is likely random contamination introduced during the pouring of the plates on the bench in a busy lab. Contaminants can be very easily and effectively removed using a flame-heated stainless steel spatula to scoop out the offending matter. This is extremely effective, provided the contaminant is not touching a plate wall (in which case it is impossible to aseptically remove it), and greatly extends the number of usable plates. Contaminants on the bottom of the plate (i.e., under the agar) are left in place. (c) I have found that if plates are used the day after pouring (after pre-incubating overnight as a test for sterility), contamination is of minimal concern.

11. Of course, wiping the lids again after the top agarose/cells/phage have been poured, prior to incubating the plates overnight, greatly decreases rivulets/rivers due to excess condensation. Parenthetically, I have noted that plates that are poured at the bottom of a stack tend to have droplets of condensed water on the agar surface (by virtue of resting on the ‘cool’ counter). These can be inverted, with the bottoms resting on the lids rotated 45 degrees, for about 20-60 minutes allowing the excess moisture and droplets to evaporate, before incubating these plates with the rest. Alternatively, have a ‘dummy’ (empty) plate at the bottom of a stack.

Protocol For Sterile Technique

Good sterile technique is the first and most important step in insuring consistent results when employing recombinant DNA and protein expression techniques. Sterile technique refers to procedures by which cultures may be manipulated without infecting the worker or contaminating the cultures or the laboratory environment.

Because contaminating bacteria are ubiquitous and are found on fingertips, bench tops, etc., it is important to minimize contact with these contaminating surfaces. When students are working with the inoculation loops and agar plates, you should stress that the round circle at the end of the loop, the tip of the pipetter, and the surface of the agar plate should not be touched or placed onto contaminating surfaces.

The flaming of lips of tubes and flasks must ALWAYS be done whenever culture liquid is to be poured from a container (e.g., pouring plates). Flaming should be routinely done when caps are removed from tubes during transfer of cultures. The purpose of flaming is not to sterilize, but to warm the tube and create warm air convection currents up and away from the opening. This “umbrella” of warm, rising air will help to prevent the entrance of dust particles upon which contaminating bacteria reside.

Petri dish lids prevent dust from falling directly onto plates but allow diffusion of air around the edges. There are no direct air currents into the plate, and to enter, dust particles would have to rise vertically more than a centimeter. This does not often occur because of the density of the particles. Whenever the lid is removed, it should be held over the plate as a shield. Do not place the lid on the bench top. Do not leave plates uncovered. Do not walk around the room with an open plate.

When working with cultures in testtubes, work as rapidly as is consistent with careful technique. Keep the tubes open a minimum amount of time. While the tubes are open, hold them at a 45 degree angle so that dust cannot fall into the open tube. Hold the tubes away from your face while transferring.

Testtubes are handled in the following manner:

  • The testtube is held in the left hand (for a right-handed person).??
  • The instrument (loop, pipet, or needle) is held in the right hand.??
  • The testtube cap is grasped by the little finger of the right hand, and removed.??
  • While continuing to hold the cap with the little finger, the tube is lightly flamed and the instrument is manipulated appropriately, and withdrawn.??
  • The cap is replaced on the testtube and the testtube is put back into the rack.

Label all cultures with the name or number of the organism, and your name.??

Always clean all work areas (your bench, balance area, sink area, gel area, etc.) thoroughly before leaving the laboratory! The last step before leaving the lab is to wash your hands thoroughly.

These are guidelines. You may find a set of techniques that best suite your working style. This is fine as long as you adhere to the basic concepts of good sterile technique.

Aspetic Technique

Aseptic techniques used by Cell Culture specialists in handling products from and/or mammalian cells.


To describe aseptic techniques used by Cell Culture specialists in handling products from and/or mammalian cells.


Protect eyes, mucous membranes, open cuts and wounds from contact with biohazard material.????Use gloves, goggles, mask, and protective clothing as necessary.


Laminar flow or biological safety hood as appropriate to the hazardous nature of the project.


1.??????????Wipes, lint-free.

2.??????????Disinfectant, or quaternary ammonium.

3.??????????Alcohol. 70% ethanol.

4.??????????Pipets, sterile, of appropriate size.

5.??????????Pipet-Aid, or equivalent.

6.??????????Biohazard waste container.


1.??????????Carry out all culturing operation is a laminar flow hood.

2.??????????Disinfect all surfaces prior to use with a disinfectant solution.

3.??????????Swab down the working surface liberally with 70% ethanol.

4.??????????Periodically spread a solution of 70% ethanol over the exterior of gloves to minimize contamination.????Replace them if torn.

5.??????????In case of any spill, spread a solution of 70% alcohol and swab immediately with non-linting wipes.

6.??????????Discard gloves after use and do not wear them when entering any other lab area.

7.??????????Bring into the work area only those items needed for a particular procedure.

8.??????????Leave a wide clear space in the center of the hood (not just the front edge) to work on.????Do not clutter the area to prevent blockage of proper air flow and to minimize turbulence.

9.??????????Swab with 70% alcohol all glassware (medium bottles, beakers, etc.) before placing them inside the hood.

10.????Arrange the work area to have easy access to all of it without having to reach over one item to get at another (especially over an open bottle or flask).

11.????Use sterile wrapped pipets and discard them after use into a biohazard waste container.

12.????Check that the wrapping of the sterile pipet is not broken or damaged.

13.????Inspect the vessels to be used:

o????????T-flask – Must be free from visible contamination or breakage, or lack container identification. The plastic covering the flask must be intact.

o????????Bottles – Check for cracks, expiration dates.

o????????Spinner flasks – Check for cracks, expiration dates, and proper assembly.

14.????Discard any biohazard or contaminated material immediately.

15.????Never perform mouth pipetting.????Pipettor must be used.

16.????When handling sterile containers with caps or lids, place the cap on its side if it must be laid on the work surface.

17.????Make sure not to touch the tip of the pipette to the rim of any flask or sterile bottle.

18.????Clean the work area when finished by wiping with 70% alcohol.

40X Tae Electrophoresis Buffer

This protocol makes 1000ml of 40X TAE.


193.8 gm Tris??(Final Concentration 1.6M)

108.9gm NaAc.3H20??(Final conc 0.8M)

14.9gm EDTA Na2.2H20??(Final Conc 40mM)


Dissolve the above reagents in?? ~ 700ml dH20

Adjust pH to 7.6 with HCl or Acetic acid

Bring volume to 1000ml

Yepd Plates

  1. Per 1L of YEPD liquid, add 20g Agar.
  2. Autoclave, cool, pour plates.

Yepd Liquid

  • Mix in bucket:
For 1L For 4L
Bacto Peptone 10g 40g
Extract of Yeast, granulated 20g 80g
Glucose 20g 80g
dH2O up to 1L 4L
  • Autoclave in 500 ml bottles

Te Buffer

Per 1L:

  • 10ml 1M Tris pH 7.5 or 8.0
  • 2ml 0.5M EDTA pH 8.0
  • dH20

Mix, pH to desired pH with HCl or NaOH, autoclave in 100ml bottles.

1M Tris Base

  1. Stir 1L ddH2O
  2. Add 121.4g Tris
  3. pH to desired pH with concentrated HCl
  4. Autoclave in 100ml bottles