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A protocol for cleaning and reusing the large 25 x 25 cm plates (by Greg Stuart)
We regularly reuse our large 25x25 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.