Jun 08, 2022
Preparation of agarose pads suitable for viewing filamentous cyanobacterial microbial communities using time-lapse imaging
- Jerko Rosko1,
- Mary Coates1,
- Kelsey Cremin1,
- Christian Zerfass2,
- Orkun Soyer1
- 1University of Warwick;
- 2Jena University
Protocol Citation: Jerko Rosko, Mary Coates, Kelsey Cremin, Christian Zerfass, Orkun Soyer 2022. Preparation of agarose pads suitable for viewing filamentous cyanobacterial microbial communities using time-lapse imaging. protocols.io https://dx.doi.org/10.17504/protocols.io.81wgb65p1lpk/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it’s working
Created: April 29, 2022
Last Modified: June 08, 2022
Protocol Integer ID: 61681
Keywords: time-lapse imaging, agarose pads, filamentous cyanobacterial microbial communities
Abstract
This protocol is for viewing cultures on top of agar pads, e.g filamentous cyanobacterial microbial community (from a freshwater environment) that form biofilms and granular structures, using time-lapse imaging.
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Materials
Materials for STEP 1:Agarose pad medium
- Gelling agent: MacroSieve Low melt agarose (Flowgen Cat No H15621)
- Agarose percentage: 2%
- Medium volume: 50 mL
STEP 1: Agarose pad medium (makes approx. 50 pads)
STEP 1: Agarose pad medium (makes approx. 50 pads)
Note
Theoretically any medium solution can be used to make agarose pads. Here, we describe 2% agarose pads made with BG11+ (DSMZ Medium 1593) Note: we make the ‘BG11-Mix’ as individual components.
Into a 100 mL Duran bottle, add 1 g agarose and make up to 47.35 mL with DW.
Autoclave at 120 °C for 20 – 30 minutes. Remove at 80 °C .
Note
For accuracy, you can weigh the bottle before and after autoclaving to calculate any evaporation losses.
After autoclaving, gentle swirl the bottle to ensure the agarose is fully dissolved.
Once removed from the autoclave, keep in a hot water bath (55 °C ) to prevent agarose from setting.
Working in a sterile environment e.g Microbiological safety cabinet (MSC), add the following sterile BG11+ components:
| A | B | |
| BG11+ Top Table mix (salts) | 2 mL | |
| Trace metals | 50 μl | |
| NaNO3 solution (300 gL-1) | 250 μl | |
| BG11-Mix components | 3 x 100uL |
At this point, there are two optional routes to take:
OPTION 116 steps
Optional route 1
For agarose pads (Step 8, below), only 10 mL medium is required per session, thus, it is best to aliquot the media at this stage to avoid repeated solidification/melting cycles of the medium.
Using sterile glass bijou bottles with screw cap lids, aliquot 9.995 mL medium per bijou.
Note
These can be stored, solidified, in the cupboard until needed.
Before using any aliquot, add 5 µL vitamin B12 (or your preferred vitamin[s]) and swirl to mix.
Note
To melt solidified medium, microwave at 800w for 00:00:30 and swirl to mix. Leave to cool slightly before adding any vitamins.
Pipetting agarose pads (Makes 10 Pads)
Pipetting agarose pads (Makes 10 Pads)
1h
1h
Sterilise 20 22-mm2 cover glass slides with 70% Ethanol inside a MSC, allowing time for drying (~01:00:00 ).
1h
Keep the agarose solution at 55 °C using a water bath before moving into the MSC.
Place five glass cover slips in each if 2 x 90 mm petri dish.
Working on one glass cover slip at a time, pipette 800 mL of cooling agarose solution evenly across the cover slip (as the agarose does not spread well when sandwiched, try to reach as far into the corners without unevenly distributing the gel). Quickly remove any air bubbles using the suction of the pipette.
Note
We found 800 mL to be the ideal volume for this medium but depending on your media components and how viscous your medium is, you may want to adjust this, anywhere up to 1 mL.
Gently lower another cover slip on top of the agarose solution, to sandwich it between the two cover slips. Quickly adjust the cover slip, if required, to ensure the agarose appears level.
Note
Manipulation is not possible once the agarose sets e.g. within 00:00:30 . The warmer the medium, the easier it is to achieve an even distribution, with no adjustments necessary after adding the top slip.
Using a new pipette tip each time, repeat steps 11-12 for each pad.
Allow the pads to solidify, then seal the lid with parafilm and store in the cupboard until required.
Note
We had condensation issues when storing pads in the fridge.
The BG11+ pads are best used the day after making, but we found them also store well for 1-2 weeks (and even longer).
Agarose ‘pads’ in 60mm dish (makes 15) (Alternative Step 2)
Agarose ‘pads’ in 60mm dish (makes 15) (Alternative Step 2)
2h
2h
| A | B | |
| Petri dish (Falcon, Cat 353002) | 60 mm | |
| Medium volume | 7 mL |
Pipette 7 mL medium per dish, leave to dry with lids on in the MSC for 01:00:00 , then a further 01:00:00 with lids off.
2h
Parafilm close the dishes and store in the cupboard for use the next day.
Note
Pads can be used on the same day, but you may get ‘drift’ when imaging due to the continued setting of the agarose.
STEP 3: Microscopy on the agarose pads
STEP 3: Microscopy on the agarose pads
When required for use with microbial cultures, the top cover slip on an agarose pad should be removed from the selected pad. The pad can then be cut into smaller pieces (if required) or the whole pad placed into a 60mm dish.
Note
You can use the top coverslip to help release the pad from the bottom coverslip and slide it into the petri dish.
Add bacteria culture and start imaging.