Multiwell Plates, Blocks, Reservoirs, Seals

Microbial culture in blocks

• Shaking cultures of ⅓ the well volume can be regarded as the maximum that prevents cross-well contamination, at least at 800 rpm. At maximum agitation, the liquid in a well ⅓ full may reach double its static height, ⅔ of well, leaving the top ⅓ of the well as a dry buffer zone against cross-contamination. Adhesive seals are used nevertheless, but better not to rely solely on it.
• Culturing 2 mL volumes allows later centrifugation/minipreps in 2 mL microcentrifuge tubes, compatible with the safe shaking well volume of 24-well blocks.
• Square wells with pyramidal "V-bottoms" are best for agitation/aeration than round wells and U-bottoms. Analytical Sales sells the following:
  
  
• 24-well block
  • ≈10 mL well volume to the top.
  • 3.3 mL max shaking culture volume.
  • Other types/lids
• 48-well block

  • 5 mL well volume

  • 1.7 mL max shaking culture volume.

• 96-well block
  • ≈2 mL well volume to the top.
  • 0.7 mL max shaking culture volume.
  • Other types

Adhesive Seals

Ideally, seals minimize cross-contamination and evaporation and allow uniform O₂/CO₂ exchange for all wells, unlike plate lids which favor exchange for wells near the plate edges.
Use of brayer ("plate roller") assures a secure uniform seal around all wells.

• AeraSeal™ (Excel Scientific) ≈ Breathe-EASIER (Diversified Biotech)
  • Maximal aeration/gas exchange; at the expense of still considerable evaporation over longer culture time. Best used with ≥0.5 mL cultures. Ideal for non-experimental growth (DNA/strain construction).
  • Material: 140 µm thick hydrophobic porous rayon with medical-grade adhesive
  • Sterile, tip-pierceable, -20°C–80°C.
  • Gas exchange rates of O₂, CO₂, and H₂O: ~4.2 kg/m²/day
  • Directions: partially peel off one side of the main bottom paper, and holding via the protected tabs, align and adhere the exposed adhesive to the plate; then adhere the remainder as you fully peel off the paper across its length, ensuring it adheres flat with no bubbles or creases. Complete the seal by rolling a brayer firmly on top. Removing the bottom paper for the tabs is not necessary; the tabs allow the film to be easily peeled on/off later.
  • For normal cloning of plasmids, it is effective to use as many wells as needed, and then save blocks with the AeraSeal first used still affixed, and later use any remaining unused wells with the same seal. This is best done by never fully removing the AeraSeal at one end, thus preserving its original alignment with the wells. The adhesive is strong enough for several rounds of peeling and reaffixing with a brayer.
• Breathe-Easy™ (not recommended)
  • Best for evaporation protection, but slower bacterial growth because of impaired gas exchange.
    • David Zong found ~½ fluorescent protein expression with Breathe-Easy compared to open wells, in 96-well assay plate. Wells under a small crease in the film grew 2-fold more than properly sealed wells.
  • Material: polyurethane with acrylic adhesive.
  • "Permeable to oxygen, carbon dioxide, and water vapor"; sterile, tip-pierceable, -80°C–100°C
  • Gas permeability: O₂ 553.5 cm³/100 in²/day; CO₂ 3781 cm³/100 in²/day; H₂O 0.7 kg/m²/day
  • Directions: partially peel off the the left side edge of the bottom paper, and holding via the left protected tab, align and adhere the edge of the exposed adhesive to the plate. 
    Place a flat, straight item on the adhered edge and use it to uniformly push down the remainder of the seal's adhesive bottom while you slowly fully peel off the back paper across the seal's length, ensuring it adheres flat with no bubbles or creases. It is essential that the seal is completely flat, or else wells with bubbles/creases will have greater oxygenation and non-uniform growth relative to other wells. Complete the seal by rolling a brayer firmly on top, and only then remove the top paper to expose the membrane.
• Silicone and santoprene lids can be purchased

Cleaning Procedure

1. Add bleach to wells containing culture, to a rough final 10%, and add 10% bleach to empty, contaminated wells, wetting the well walls. Let sit 20–30 min before draining and rinsing in tap water, scrubbing out any adhering material.
   Blocks should appear spotless when held up to light; no gunk.
2. Lastly, fill the block with tap water and leave on your bench or by the used blocks bin, allowing the water to extract off residual bleach. Eventually, drain these and place in the used blocks bin.
3. Sterilization — Fill used blocks from bin with DI-water to the brim, place in one–two layers in clean autoclave bins, and autoclave 45 min on the liquid cycle.
   As soon as the cycle is over, while the contents is still hot, the drain in block into a sink and fling the residual water out. Place blocks in an ethanol-wiped autoclave bin standing on their side to dry off over 12 hr.
4. Storage — Once dry, stack blocks of similar kinds, covering the topmost block with foil. Invert the stacks and place inverted in the blocks cabinet to protect the wells from dust. This also makes the block layout apparent since the bottoms are up.
   
   Typical culturing with antibiotic selection and fast-growing E. coli makes more stringent cleaning and sterility practices not useful.
   If useful, one can better ensure destruction of organic compounds in blocks by submerging them in a large beaker of 10% bleach to fill wells to the brim, letting the bleach act for 20 min, and proceeding on with rinsing and sterilization as above. Where environmental contamination can be an issue, blocks can be covered individually with foil and dry-autoclaved 15 min.