Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
July 21, 2009
rev. 0.1
Overview:




[Protocol 1.0]
[Protocol 1.1]
[Protocol 1.2]
[Protocol 1.3]


[Protocol 2.0] Cleaning a Previously Unused MEA
[Protocol 2.1] MEA Sterilization with ETOH and UV light


[Protocol 3.0] PEI plus Laminin MEA Coating
[Protocol 3.1] Prepare Culture Medium

[Protocol 4.0] Cortical Cell Thaw and Culture

[Protocol 5.0] Medium Exchange
Clean and sterilize glassware
Clean and sterilize screw caps
Prepare and sterilize ultrapure water
Autoclave FEP-Sealed MEA Cover and Pipet Tips
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
July 21, 2009
rev. 0.1
Revision History:
Release Level
Date
July 14, 2009
July 21, 2009
Initial Draft
Reviewed and Accepted Protocol
Version
0.0
0.1
References:
[1]
R. I. Freshney, Culture of Animal Cells: A Manual of Basic Technique, 5th Edition. Wilmington, DE:
Wiley-Liss, 2005.
[2]
Multichannel Systems, “Microelectrode Array (MEA) User Manual,” Innovations in
Electrophysiology: Multi Channel Systems. [Online]. Available:
http://www.multichannelsystems.com/fileadmin/user_upload/Manuals/MEA_Manual.pdf.
[Accessed: May 19, 2009].
[3]
Multichannel Systems, “Microelectrode Array (MEA) Cleaning Quick Guide,” Innovations in
Electrophysiology: Multi Channel Systems. [Online]. Available:
http://www.multichannelsystems.com/fileadmin/user_upload/Manuals/MEACleaningQuickGuide.pdf. [Accessed: July 13, 2009].
[4]
S. M. Potter and T. B. DeMarse, “A new approach to neural cell culture for long-term studies,”
Journal of Neuroscience Methods, vol. 110, no. 1-2, pp. 17-24, 2001.
[5]
Brain Bits, “Frozen Neuron Thaw and Culture Protocol for Vials of One Million Cells,” Frozen Cells
1 Million Protocol: Brain Bits LLC. [Online]. Available:
http://www.brainbitsllc.com/frozencells1millionprotocol.aspx. [Accessed: July 13, 2009].
[6]
G. J. Brewer, J. R. Torricelli, E. K. Evege, and P. J. Prince, “Optimized Survival of Hippocampal
Neurons in B27 Supplemented Neurobasal,” Journal of Neuroscience Research, vol. 35, no. 5, pp.
567-576, 1993.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
1.0
July 21, 2009
rev. 0.1
Clean and Sterilize Glassware
The following protocol has been reproduced from [1], Protocol 11.1 Preparation and Sterilization of
Glassware, page 147.
Materials:
Material
Graduated Autoclavable
Screw-Cap Bottles
Disinfectant
Detergent
Soaking Baths
Bottle Brushes
Stainless Steel Baskets
Aluminum Foil
Sterility Indicators
Sterile-indicating tape or
tabs (Bennett)
Sterilizing Oven
Description
Graduated glass borosilicate (e.g. Pyrex) or autoclavable plastic
(polycarbonate) screw-cap bottles (ensure sufficient head space for later
additions), 500mL.
Hypochlorite, 300ppm available chlorine, minimum, when diluted in
detergent (e.g. Precept tablets, Clorox or Chloros; see Appendix II)
e.g., 7XTM or Decon®
To collect washed and rinsed glassware for drying
Alpha Medical for dry heat or steam sterilization; Thermalog indicators for
steam sterilization only
This is different from the sterile-indicating tape used in autoclaves, as the
sterilizing temperature is higher in an oven. Most autoclave tapes tend to
char and release traces of volatile material from the adhesive, which can
leave a deposit on the oven or even the glassware.
Fan assisted, capable of reaching 160°C, and preferably with recording
thermometer and flexible probe.
Protocol:
Collection and washing of glassware:
1. Immediately after use, collect glassware into detergent containing disinfectant. It is important
that glassware does not dry before soaking, or cleaning will be much more difficult.
2. Soak overnight in detergent.
3. Rinsing
a. Brush glassware by hand or with a mechanized bottle brush the following morning, and
rinse thoroughly in four complete changes of tap water followed by three changes of
deionized water. A sink-rinsing spray is a useful accessory; otherwise bottles must be
emptied and filled completely each time. Clipping bottles in a basket will help to speed
up this stage.
b. Machine rinses should be done without detergent. If done on a spigot header, this can
be reduced to two rinses with tap water and one with deionized or reverse-osmosis
water.
4. After rinsing thoroughly, invert bottles, etc., in stainless steel wire baskets and dry upside down.
5. Cap bottles with aluminum foil when cool, and store.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
July 21, 2009
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Sterilization of Glassware:
1. Attach a small square of sterile-indicating tape or other indicator label to glassware, and date.
2. Place glassware in an oven with fan-circulated air and temperature set to 160°C.
3. To ensure that the center of the load reaches 160°C:
a. Place a sterility indicator in a bottle or typical item in the middle of the load.
b. If using a recording thermometer, place the sensor in a bottle or typical item in the
middle of the load.
c. Do not pack the load too tightly; leave room for circulation of hot air.
4. Close the oven, check that the temperature returns to 160°C, seal the oven with a strip of tape
with the time recorded on it (or use automatic locking and recorder), and leave for 1h.
5. After 1h, switch off the oven and allow it to cool with the door closed. It is convenient to put
the oven on an automatic timer so that it can be left to switch off on its own overnight and be
accessed in the morning. The precaution allows for cooling in a sterile environment and also
minimizes the heat generated during the day, when it is hardest to deal with.
6. Use glassware within 24-48h.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
1.1
July 21, 2009
rev. 0.1
Clean and Sterilize Screw Caps
The following protocol has been reproduced from [1], Protocol 11.3 Preparation and Sterilization of
Screw Caps, page 153.
Materials:
Material
Screw-Caps
Disinfectant
Detergent
Soaking Baths
Stainless Steel Baskets
Sterility Indicators
Glass Petri Dishes
Autoclavable Plastic Film
or Paper Sterilization
Bags
Sterile-indicating
autoclave tape
Autoclave
Description
Screw caps to fit.
Hypochlorite, 300ppm available chlorine (e.g. Precept tablets)
(e.g., 7XTM or Decon®)
(to collect caps for washing and drying)
(e.g., see also Appendix II)
(for packaging)
(see Appendix II)
Autoclave, with recording thermometer with flexible probe that can be
inserted in load.
Protocol:
Collection and washing:
Metal or phenolic caps with liners:
1. Soak 30 min (maximum) in detergent.
2. Rinse thoroughly for 2h. (Make sure all caps are submerged.) Liners should be removed and
replaced after rinsing, which may be carried out in either of two ways:
a. In a beaker (or pail) with running tap water led by a tube to the bottom. Stir the caps by
hand every 15 min.
b. In a basket or, better, in a pipette-washing attachment. Rinse in an automatic washing
machine, but do not use detergent in the machine.
Polypropylene caps:
These may be washed and rinsed by hand as just described (extending the detergent soak if necessary).
Because these caps may float, they must be weighted down during soaking and rinsing. For automatic
washers, after soaking in detergent, use pipette-washing attachment and normal cycle without machine
detergent.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
July 21, 2009
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Stoppers:
Shrouded caps are preferred to stoppers, but if the latter are required, use silicone or heavy metal-free
white rubber stoppers in preference to those made of natural rubber. Wash and sanitize as for caps.
(There will be no problem with floatation in washing and rinsing.)
Sterilization:
3. Place caps in a glass Petri dish with the open side down.
4. Wrap Petri dish containing caps in cartridge paper or steam-permeable nylon film, and seal with
autoclave tape (Fig 11.8 [pg 153]).
5. Prepare similar package with sterility indicator enclosed and insert in the middle of the load.
6. Autoclave for 20 min at 121°C and 100kPa (15 p.s.i.) (Fig 11.9 [pg 153]).
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
1.2
July 21, 2009
rev. 0.1
Preparation and Sterilization of Ultrapure Water (UPW)
The following protocol has been reproduced from [1], Protocol 11.5 Preparation and Sterilization of
Ultrapure Water (UPW), page 156.
Materials (Nonsterile):
Material
Graduated Autoclavable
Screw-Cap Bottles
Screw-Caps
Sterility Indicator Strips
Sterile Indicating Tape
Marker Pen
Water Purification
Equipment
Autoclave
Log Book
Description
Graduated glass borosilicate (e.g. Pyrex) or autoclavable plastic
(polycarbonate) screw-cap bottles (ensure sufficient head space for later
additions), 500mL.
Screw caps to fit.
Marker pen or preprinted labels.
Record of preparation and sterilization.
Standard Protocol:
1. Label bottles with date, contents, and batch number.
2. Early in the morning after overnight recycling, run about 50mL water to waste from purifier,
check conductivity (or resistivity) and total organic carbon (TOC) on respective meters, and
enter in log book.
3. If water is within specified limits (resistivity ≥10MΩ-cm at 25°C, TOC ≤10ppb), collect ultrapure
water directly into labeled bottles.
4. Fill to the specified mark, e.g., 430-450mL if to be used for diluting 10x concentrated medium
(see Protocol 11.8). If to be autoclaved open, add 10% extra.
5. Place sterility indicator in one bottle (to be discarded when checked after autoclaving).
6. Seal bottles with screw caps. If using soda glass, i.e., if the bottle is liable to breakage, then
leave caps slack.
7. Place bottles in autoclave with bottle containing sterility indicator in center of load.
8. Close autoclave and check settings: 121°C, 100kPa (15lb/in.2, 1bar), for 20 min with postvac
deselected.
9. Start sterilization cycle.
10. On completion of cycle, check printout to confirm that correct conditions have been attained for
the correct duration and enter in log book.
11. Allow load to cool below 50°C.
12. Open autoclave and retrieve bottles. If caps were slack during autoclaving, tighten when bottles
reach room temperature.
Note: Sealing bottles, which have been open during autoclaving, when they are still warm can
cause the liner in some caps to be drawn into the bottle as it cools and the contents contract.
Also, there will be a rapid intake of air into the bottle when it is first opened, and this can cause
contamination. Bottles that have been open during autoclaving should be allowed to cool to
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
July 21, 2009
rev. 0.1
room temperature in a sterile atmosphere, e.g., in the autoclave or in horizontal laminar flow,
before the caps are tightened.
Check sterility indicator to confirm that sterilization conditions have been achieved and enter in
log book.
13. Place bottles in short-term storage at room temperature.
14. Replenish culture room stocks as required, confirming by examination of sterile-indicating tape
that the bottles have been through sterilization cycle.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
1.3
July 21, 2009
rev. 0.1
Autoclave FEP-Sealed MEA Cover and Pipet Tips
The following protocol has been adapted from [1].
Materials:
Material
MEA Covers
Small Pipet Tips
Large Pipet Tips
Description
Hard plastic ring with o-ring seals and FEP cover
200µL Eppendorf Pipet Tips in Autoclavable Container
5000µL Eppendorf Pipet Tips in Autoclavable Container
Protocol:
1. Place MEA covers in a glass Petri dish with the open side down.
2. Wrap Petri dish containing covers in cartridge paper or steam-permeable nylon film, and seal
with autoclave tape.
3. Prepare similar package with containers of pipet tips enclosed.
4. Prepare similar package with sterility indicator enclosed and insert in the middle of the load.
5. Autoclave for 20 min at 121°C and 100kPa (15 p.s.i.).
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
2.0
July 21, 2009
rev. 0.1
Cleaning a Previously Unused MEA
The following protocol has been adapted from [2] and [3].
Materials:
Material
MEA
Tissue Culture Water
Terg-A-Zyme
Description
Autoclaved ultrapure water.
(Sigma, order number Z273287)
Protocol:
1.
2.
3.
4.
5.
Prepare a 1% solution of Terg-A-Zyme, diluted in tissue culture water.
Place MEA in this solution in laminar flow hood.
Apply gentle shaking/rocking (if possible).
Allow to soak overnight at room temperature.
Rinse thoroughly with tissue culture water.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
2.1
July 21, 2009
rev. 0.1
MEA Sterilization via Ethanol and Ultraviolet Light
The following protocol has been adapted from [2].
Materials:
Material
MEA
Tissue Culture Water
Ethanol
Description
Autoclaved ultrapure water.
Protocol:
1. Prepare a 70% solution of ethanol, diluted in tissue culture water.
2. Rinse MEA with this solution in laminar flow hood.
3. Allow MEA to air dry over night in laminar flow hood with UV light turned on.
Note: Alternatively, MEAs can be sterilized by autoclave (134°C for 3 min), dry heat (121°C for
15 min), or with hot water (90°C for 1 min).
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
3.0
July 21, 2009
rev. 0.1
PEI plus Laminin MEA Coating
The following protocol has been adapted from [2], “5.4.2 Coating with Polyethyleneimine (PEI) (plus
Laminin),” page 20, and [4], “4.2 MEA Preparation.”
Materials:
Material
MEA
Tissue Culture Water
Poly(ethyleneimine)
Solution (PEI)
Boric Acic, Crystalline
Borax (sodium
tetraborate)
1N HCl
Laminin, 1mg/mL
Description
Cleaned and sterilized according to protocol.
Autoclaved ultrapure water.
(Sigma-Aldrich, Inc., P3143)
(Fisher Scientific, A73-500)
(Sigma-Aldrich, Inc., B0127)
(Sigma-Aldrich, Inc., L2020)
Borate Buffer:




3.10g boric acid
4.75g borax
Dissolve in 1L distilled water.
Adjust pH to 8.4 with 1N HCl.
PEI Stock Solution:

0.05-0.1% PEI dissolved in borate buffer.
Laminin Solution:

20µg/mL laminin in plating medium (20µL of 1mg/mL laminin was diluted in 1mL of medium
according to [4].)
Protocol:
Note: It is necessary to thoroughly rinse off unbound PEI from the plates before use, as dried PEI is
toxic.
1.
2.
3.
4.
5.
6.
Pipette 500µL PEI solution onto MEA. The recording field should be completely covered.
Incubate at room temperature for 1h, or at 4°C over night.
Remove PEI solution and thoroughly rinse 4x with tissue culture water.
Air-dry the MEA in the laminar flow hood with the UV light turned on.
Allow UV light to sterilize MEA for at least 1h after coating.
Apply 10µL droplet of sterile laminin solution onto the center of the MEA, covering the
electrode region, and incubate for 30 min.
7. Aspirate, do not rinse, and directly seed your cells.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
3.1
Prepare Culture Medium
The following protocol has been adapted from [5] and [6].
Materials:
Material
Neurobasal Culture
Medium
B-27
GlutaMAX-I
Description
Neurobasal Medium (1x) Liquid, 500mL, Invitrogen
B-27 Serum Free Supplement (50x) Liquid, 10mL, Invitrogen
GlutaMAX-I Supplement, 200mM supplied in 0.85% NaCl.
Protocol:
1. To Neurobasal medium add 0.5mM GlutaMAX-I supplement.
𝐶𝑖 ∗ 𝑉𝑖 = 𝐶𝑓 ∗ 𝑉𝑓
200𝑚𝑀 ∗ 𝑉𝑖 = 0.5𝑚𝑀 ∗ 500𝑚𝐿
𝑉𝑖 =
(0.5𝑚𝑀 ∗ 500𝑚𝐿)
200𝑚𝑀
𝑉𝑖 = 1.25𝑚𝐿
2. Add 10mL of B-27 (2mL/100mL) to Neurobasal medium.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
4.0
July 21, 2009
rev. 0.1
Cortical Cell Thaw and Culture
The following protocol has been adapted from [4] and [5].
Materials:
Material
MEA
Cortical Cells
Tissue Culture Water
Glass Coverslips
Medium
15mL Sterile Polystyrene
or PET Centrifuge Tube
Ethanol
Water bath
Hemacytometer
Trypan Blue
Description
Cleaned and sterilized, then coated with PEI plus laminin according to
protocol.
Frozen cells to be cultured.
Autoclaved ultrapure water.
Neurobasal, 50x B27, 0.5mM glutamine or Glutamax, all Invitrogen
(Corning #430055)
Diluted to 70% in tissue culture water.
37°C
(VWR #15170-079)
(Sigma #T8154)
Growth Medium:


Neurobasal, 2% B-27, 0.5mM Glutamax or complete NbActiv4.
Bring to room temperature.
Protocol:
1. Thaw one tube at a time for a minimum amount of time in 37°C water bath. (Target 3 min.)
Remove immediately when thawed.
Note: Do not vortex or shake vigorously. Wipe outside of cryovial with 70% ETOH.
2. Transfer 0.21mL to a 50mL centrifuge tube without creating bubbles.
3. Add 1.91mL room temperature medium dropwise over a period of 30 seconds. Mix gently as
you add medium. Gently mix by inversion two times.
4. Immediately plate 0.2mL/cm2 substrate (range 0.15-0.5mL/cm2) or 2mL per dish containing 3
precoated slips. Gently mix cell suspension every 2 minutes in between aliquots.
5. Incubate plated cells 37°C, 5% CO2, 9% O2, (ambient oxygen will work also). Also place more
medium in incubator with loose cap to equilibrate pH for step 7.
6. Mix leftover cells from step 4 and aliquot 30µL of cells into 30µL of trypan blue.
7. After 0.7-3h incubation, remove culture from incubator for medium exchange. Fill pipet with
fresh medium equilibrated in 37°C, 5% CO2, 0.2 or 0.4mL fresh medium/ cm2 (for 35mm dish use
2.0mL). Drain and aspirate medium over cells and immediately add fresh medium. Be careful
not to let cells dry out. Be careful to add medium gently from side so that adherent cells are not
disturbed. Quickly observe phase bright adherent cells with 10 or 20x objective mag to ensure
attachment to substrate.
8. After four days in culture or longer, cells are ready for your use.
9. One-half medium should be exchanged once or twice a week with equilibrated medium.
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10. Measure viability at 4 to 5 DIV. Exchange medium with warm Hibernate E/B27/gIn. Count live,
phase-bright cells with processes. Small, bright cells are dead or apoptosing.
Live-Dead Fluorescence Assay:
Note: Viability assay: 𝑣𝑖𝑎𝑏𝑖𝑙𝑖𝑡𝑦 =
𝑔𝑟𝑒𝑒𝑛 𝑐𝑒𝑙𝑙𝑠 𝑝𝑒𝑟 𝑢𝑛𝑖𝑡 𝑎𝑟𝑒𝑎
𝑡𝑜𝑡𝑎𝑙 𝑐𝑒𝑙𝑙𝑠 𝑝𝑙𝑎𝑡𝑒𝑑 𝑝𝑒𝑟 𝑢𝑛𝑖𝑡 𝑎𝑟𝑒𝑎
or 𝑠𝑢𝑟𝑣𝑖𝑣𝑎𝑙 =
𝑔𝑟𝑒𝑒𝑛 𝑐𝑒𝑙𝑙𝑠
.
𝑔𝑟𝑒𝑒𝑛+𝑟𝑒𝑑 𝑐𝑒𝑙𝑙𝑠
1. Rinse 2x with PBS of HBSS, 0.4mL/cm2 of substrate.
2. From an acetone stock of 15mg/mL fluorescein diacetate (Sigma #F7378), add 15µL (1:100
dilution of the stock into 1.5mL HBSS). From an aqueous stock of 4.6mg/mL propidium iodide
(Sigma #P4170), add 15µL of the stock into the same 1.5mL HBSS (1:100 dilution). Add 40µL of
that dilution to each well with 0.4mL HBSS (further 1:10 dilution).
3. After about 1 min, count using Nikon B1A, G1B filters or other blue excitation appropriate for
fluorescein fluorescence. Green cells are live. Small red nuclear 3 stain indicates a dead cell.
If desired, fix and stain with 0.25% Commassie blue R in ethanol/HAc/water (45/10/45), 1 min, rinse
with 10% HAc, aspirate and dry.
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Department of Electrical and Computer Engineering
Neurobiology Engineering Laboratory
5.0
July 21, 2009
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Medium Exchange
The following protocol has been adapted from [4] and [5].
Materials:
Material
Complete Medium
Description
Protocol:
1. Place medium in incubator with loose cap too equilibrate pH and temperature for medium
exchange.
2. Incubate for 1h.
3. Wipe down laminar flow hood with 70% ETOH.
4. Sterilize laminar flow hood via UV light for 20 min.
5. Remove existing medium via suction by tilting MEA.
Note: Do not touch the MEA or suction near the electrode area!
Note: Do not allow the cells to dry out!
6. Replace with fresh, equilibrated medium immediately. Fill pipet with 0.4mL/cm2 (for 35mm dish
use 2.0mL fresh, equilibrated medium).
7. Replace FEP culture chamber lid and return culture to incubator.
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