Bridges Lab Protocols - User contributions [en]

SOP - Animal Anesthetics

2019-01-08T18:07:08Z

Lgunder:

[[ Category: SOP ]]
[[ Category: Lab Safety]]

==Description==
This standard operating procedure outlines the handling and use of animal anesthetics including: isoflurane, halothane, enflurane and ether. Review this document and supply the information required in order to make it specific to your laboratory. In accordance with this document, laboratories should use appropriate controls and personal protective equipment when handling animal anesthetics.

==Procedure Location==
The use of animal anesthetics must be performed in an area with good ventilation and controls to capture and exhaust waste anesthetic gases.

==Potential Hazards==
Anesthetic gas and vapor that leaks during medical or research procedures are considered waste anesthetic gases (WAGs). University faculty, staff and students should be aware of the potential risks of WAGs and be advised to take appropriate precautions to reduce exposures. Workers acutely exposed to excessive amounts of anesthetic gas can experience symptoms of drowsiness, headache, nausea, poor judgment and loss of coordination. Chronic symptoms of over-exposure can include liver, kidney and reproductive effects. Safety precautions include the use of an approved gas scavenging system, or using the agent inside a certified chemical fume hood.

The use of ether is not recommended because it is flammable and a mutagen. Be certain that there are no ignition sources present when handling ether. There are restrictions concerning the use of ether with animals. Contact OSEH at (734) 647-1143 concerning the use of ether.

==Engineering Controls==
Anesthetics should not be handled on the bench top without special ventilation or a scavenging system. Anesthetic gas filtering cartridges, snorkel exhaust, fume hoods or other scavenging systems must be used. ULAM provides ventilated procedure rooms designed for use of anesthetic gases in many areas. Fume hoods provide the best protection against exposure to anesthetics in the laboratory and are the preferred ventilation control device when handling greater than 100 cc outside of the original container. Always handle large quantities of ethyl ether in a fume hood due to the flammable nature of the material. If your research does not permit the handling of large quantities of ethyl ether in your fume hood, contact OSEH to review the adequacy of all special ventilation.

Liquid anesthetics administered with a vaporizer must be scavenged. When used properly, vaporizers equipped with activated charcoal canisters (e.g. F/Air) are effective in removing halogenated waste gases. The F/Air canister containing activated charcoal will absorb waste anesthetics for about 12 hours. Note: F/Air Canisters only absorb halogenated anesthetics (e.g. isoflurane, halothane). Immediately before using any anesthesia machine, the F/Air canister should be removed and weighed to evaluate the remaining absorption capacity. The weight should be recorded and dated on the side of the canister. Immediately following the use of an anesthesia machine, the number of hours the machine was in use should be recorded next to the dated weight information.

Canisters that exceed 12 hours of use or 50 grams of accumulated weight (whichever comes first) must be removed and placed in a sealed plastic bag and disposed of as a hazardous waste through OSEH Hazardous Materials Management (HMM) at (734) 763-4568. Thoroughly clean the induction chamber immediately after each use to avoid residual anesthetic waste release into the environment (which can continue to be released for up to three hours). Please refer to the OSEH Guideline entitled Anesthetic Gas Use for additional information on the safe use of anesthetic gases.

==Work Practice Controls==
All anesthetic agents must be clearly labeled with the correct chemical name. Handwritten labels are acceptable; chemical formulas and structural formulas are not acceptable.

Always keep the flow rate of anesthetics to the animal as low as possible during the procedure. High flow rates can increase your exposure to the anesthetic. It is also important to move the point of potential gas release as close to the exhaust system as possible to increase capture of the chemical.

Do not permit containers to remain open on the bench top. The odor thresholds for most liquid anesthetics (except for ether) are well above permissible exposure limits. If you smell the anesthetic the control procedures you are using are inadequate and must be re-evaluated.

==Personal Protective Equipment (PPE)==
Eye protection in the form of safety glasses must be worn at all times when handling anesthetic agents. Ordinary (street) prescription glasses do not provide adequate protection

Single use nitrile or latex gloves must be worn when handling anesthetic agents as well as lab coats, closed toed shoes and pants. Additional protective clothing should be worn if the possibility of skin contact is likely.

==Transportation and Storage==
Ethers form potentially explosive peroxides after exposure to air and light. Since these chemicals are packaged in an air atmosphere, peroxides can form even though the containers have not been opened. Write the date received and date opened on all containers of ether. Opened containers of ether should be discarded within 12 months of opening. Even closed containers of ether must be discarded by the expiration date through OSEH-HMM (734) 763-4568.

Halogenated liquid anesthetic agents (i.e. halothane, enflurane, isoflurane) are not flammable but do have limited shelf life. Be certain to date the chemical when it is opened and to check expiration date before use.

Always purchase the smallest quantity required for use. Ether used for anesthetic purposes should be purchased in the smallest quantity available (typically 150 cc, Fisher Scientific E136-150) due to its short (12 month) shelf life after it is opened.

==Waste Disposal==
Anesthetic agents are hazardous wastes. Contact OSEH-HMM at (734) 763-4568 for waste containers, labels, manifests, waste collection and for any questions regarding proper waste disposal. Also refer to OSEH’s Hazardous Waste webpage for more information.

==Exposures/Unintended Contact==

'''If the employee is in need of emergency medical attention, call 911 immediately.'''

Wash hands and arms with soap and water immediately following any skin contact with anesthetic agents. Flush eyes for 15 minutes following eye contact.

Contact OSEH for advice on symptoms of chemical exposure, or assistance in performing an exposure assessment.

Report all work related accidents, injuries, illnesses or exposures to WorkConnections within 24 hours by completing and submitting the Illness and Injury Report Form. Follow the directions on the WorkConnections website Forms Instructions to obtain proper medical treatment and follow-up.

Complete the OSEH Laboratory Incident and Near-Miss Report form.

==Treatment Facilities:==
*U-M Occupational Health Services -- Campus Employees
Mon-Fri 7:30 am - 4:30 pm
After hours - go to UM Hospital Emergency Dept. – Urgent Care Clinic
C380 Med Inn building
1500 East Medical Center Drive, Ann Arbor (734) 764-8021
*University Health Services -- University students (non-life threatening conditions)
Mon-Fri 8 am – 4:30 pm, Sat 9 am – 12 pm
Contact for current hours as they may vary
207 Fletcher Street, Ann Arbor (734) 764-8320
*UMHS Emergency Department -- after clinic hours or on weekends
1500 East Medical Center Drive, Ann Arbor, (734) 936-6666

Click here for additional accident and injury reporting information.

==Spill Procedure==
Ether is extremely flammable. If ether is spilled immediately assess and deactivate potential ignition sources. Be prepared for a potential fire and ensure your safety and others first.

Anticipate spills by having the appropriate clean up equipment on hand. Spill materials for anesthetic agents are designed to control the liquid portion of the spill and minimize the production of vapors. Never use paper towels on large spills of anesthetic agents because it exacerbates vapor production.

*When a spill occurs, personal safety should always come first.
*Alert and clear everyone in the immediate area where the spill occurred.

A minor (small) chemical spill is one that the laboratory staff is capable of handling safely without the assistance of safety and emergency personnel, i.e., less than 1 liter. A major/large chemical spill requires active assistance from emergency personnel.

==Additional Spill Response Steps:==

===MINOR CHEMICAL SPILL===
*Alert people in immediate area of spill.
*If spilled material is flammable, turn off ignition and heat sources. Don’t light Bunsen burners or turn on other switches.
*Open outside windows, if possible.
*Wear protective equipment, including safety goggles, gloves and long-sleeve lab coat.
*Avoid breathing vapors from spill.
*Confine spill to as small an area as possible.
*Do not wash spill down the drain.
*Use appropriate spill kits/sorbents to absorb spill. Collect contaminated materials and residues and place in container. Contact OSEH-HMM (734) 763-4568 for proper disposal.
*Clean spill area with water.

===MAJOR CHEMICAL SPILL===
*Attend to injured or contaminated persons and remove them from exposure.
*Alert people in the laboratory to evacuate.
*If spilled material is flammable, turn off ignition and heat sources. Don’t light Bunsen burners or turn on other switches.
*Call University of Michigan Division of Public Safety and Security (DPSS) at 911 immediately for assistance.
*Close doors to affected area.
*Post warnings to keep people from entering the area.
*Have person available that has knowledge of incident and laboratory to assist emergency personnel.

==Additional Spill Links:==
• www.oseh.umich.edu/pdf/chemspil.pdf
• http://www.oseh.umich.edu/emer-chemical.shtml.

Report all emergencies, suspicious activity, injuries, spills, and fires to the University of Michigan Division of Public Safety and Security (DPSS) by calling 911 or texting 377911. Register with the University of Michigan Emergency Alert System via Wolverine Access.

==Training of Personnel==
All personnel are required to complete the General Laboratory Safety Training session (BLS025w or equivalent) via OSEH’s My LINC website. Furthermore, all personnel shall read and fully adhere to this SOP when handling animal anesthetics.

==Certification==
I have read and understand the above SOP. I agree to contact my Supervisor or Lab manager if I plan to modify this procedure. Sign by logging in and typing <nowiki>* ~~~~</nowiki> in the list below:

*[[User:Reddj|Reddj]] ([[User talk:Reddj|talk]]) 14:44, 13 October 2016 (UTC)

* [[User:Iharvey|Iharvey]] ([[User talk:Iharvey|talk]]) 19:34, 10 November 2016 (UTC)

*[[User:Pfeiferl|Pfeiferl]] ([[User talk:Pfeiferl|talk]]) 18:09, 5 June 2017 (UTC)

* [[User:Snyderds|Snyderds]] ([[User talk:Snyderds|talk]]) 18:34, 7 June 2017 (UTC)
* [[User:Elhabbal|Elhabbal]] ([[User talk:Elhabbal|talk]]) 21:38, 12 June 2017 (UTC)
*[[User:Mollyec|Mollyec]] ([[User talk:Mollyec|talk]]) 14:48, 13 June 2017 (UTC)
*[[User:Lgunder|Lgunder]] ([[User talk:Lgunder|talk]]) 18:07, 8 January 2019 (UTC)

Prior Approval required – Is this procedure hazardous enough to warrant prior approval from the Laboratory Director? ☐ YES X NO

Laboratory Director - Dave Bridges Revision Date - 2016-10-13

Measuring Cross-Sectional Area (CSA)

2018-08-27T18:43:08Z

Lgunder:

== Using ImageJ ==
#Drag your image into ImageJ.
#Use the Magnifying glass. Click on the image to zoom in and right click to zoom out
#Zoom in on the scale on your image
#Use the Scrolling tool(hand) to grab and move the picture
#Set the scale with the *Straight* (line tool). Click on one end of the scale and hold shift to keep the line straight to the other end
#"Analyze"
#"Set scale"
#"Known distance:" 200 (for 20x)
#"Unit of length:" um
#"OK"
#Adjust the zoom. Start in upper left hand corner
#Use the Freehand selections tool (bean) to trace the fiber by holding down mouse press "M"
#Continue tracing and press "M" after each fiber trace
#"Results"
#"Summarize"

Measuring Cross-Sectional Area (CSA)

2018-08-27T18:41:54Z

Lgunder:

== Using ImageJ ==
#Drag your image into ImageJ.
#Use the Magnifying glass. Click on the image to zoom in and right click to zoom out
#Zoom in on the scale on your image
#Use the Scrolling tool(hand) to grab and move the picture
#Set the scale with the *Straight* (line tool). Click on one end of the scale and hold shift to keep the line straight to the other end
#Analyze
#Set scale
#Known distance: 200 (for 20x)
#Unit of length: um
#OK
#Adjust the zoom. Start in upper left hand corner
#Use the Freehand selections tool (bean) to trace the fiber by holding down mouse press M
#Continue tracing and press M after each fiber trace
#Results
#Summarize

Measuring Cross-Sectional Area (CSA)

2018-08-27T18:41:11Z

Lgunder:

== Using ImageJ ==
#Drag your image into ImageJ.
#Use the Magnifying glass. Click on the image to zoom in and right click to zoom out
#Zoom in on the scale on your image
#Use the Scrolling tool(hand) to grab and move the picture
#Set the scale with the *Straight* (line tool). Click on one end of the scale and hold shift to keep the line straight to the other end

==To Analyze and Get Results==
#Analyze
#Set scale
#Known distance: 200 (for 20x)
#Unit of length: um
#OK
#Adjust the zoom. Start in upper left hand corner
#Use the Freehand selections tool (bean) to trace the fiber by holding down mouse press M
#Continue tracing and press M after each fiber trace
#Results
#Summarize

Measuring Cross-Sectional Area (CSA)

2018-08-27T18:30:26Z

Lgunder: Created page with "== Using ImageJ == #Drag your image into ImageJ. #Use the Magnifying glass. Click on the image to zoom in and right click to zoom out #Zoom in on the scale on your image #Use..."

== Using ImageJ ==
#Drag your image into ImageJ.
#Use the Magnifying glass. Click on the image to zoom in and right click to zoom out
#Zoom in on the scale on your image
#Use the Scrolling tool(hand) to grab and move the picture
#Set the scale with the *Straight* (line tool). Click on one end of the scale and hold shift to keep the line straight to the other end

==To Analyze and Get Results==
#Analyze
#Set scale
#Known distance: 200
#Unit of length: um
#OK
#Adjust the zoom. Start in upper left hand corner
#Use the Freehand selections tool (bean) to trace the fiber by holding down mouse press M
#Continue tracing and press M after each fiber trace
#Results
#Summarize

NADH NBT Staining

2018-08-27T18:20:46Z

Lgunder: Created page with "== Tris Buffer .2M pH 7.4 == #Warm Tris buffer to 37°C in water bath. #Use hot plate to keep the buffer at 37°C, pH the buffer with HCl. (pH changes with temperature) ==Met..."

== Tris Buffer .2M pH 7.4 ==
#Warm Tris buffer to 37°C in water bath.
#Use hot plate to keep the buffer at 37°C, pH the buffer with HCl. (pH changes with temperature)

==Methods==
#20mg NBT aka Nitrotetrazolium Blue Chloride or Nitro Blue Tetrazolium
#80mg of NADH aka β-Nicotinamide adenine dinucleotide, reduced disodium salt hydrate
#100mL of Tris Buffer
#100mL fits in a staining dumpster
#Incubate for 30 minutes at 37°C

== Dehydrate and coverslip ==
#Rinse 3-4 times under running DI water
#Immerse in 50% ethanol for 2x 30 seconds
#Immerse in 70% ethanol for 2x 30 seconds
#Immerse in in 95% ethanol for 2x 30 seconds
#Immerse in in 100% ethanol for 2x 30 seconds
#Immerse in 1:1 xylene ethanol for 2 minutes for 2x 30 seconds
#Repeat twice more using fresh xylene. Check staining under the microscope
#Coverslip using Permount

Preparation of RNA Samples from Mouse Tissues

2017-12-14T20:13:37Z

Lgunder:

==Safety Information==
* [[SOP-_Phenol|SOP - Phenol]]
* [[SOP_-_Chloroform|SOP - Chloroform]]

==Materials==
*PureLink RNA Mini Kit (Invitrogen cat#12183-018A)
*Mouse Tissue (50-100 mg, about a 3mm cube)
*TRIZol (Invitrogen cat# 12183-555)
*Chloroform (in solvent cabinet)
*Label tubes, for each sample need 3 eppies, at least one of which is 2 mL and one of which is a recovery tube from the PureLink kit.
*70% Ethanol make with RNAase free water and 100% Ethanol.

==Protocol==
#Add 1 mL TRIzol reagent to each 2 mL tube.
#Cut ~50-100 mg of tissue. If tissue is in RNAlater, cut at room temperature. If tissue is frozen cut on dry ice. Weigh into a fresh 2mL eppendorf tube and keep on ice.
#Using tissue grinder, and after adding the ball bearings to every vial, homogenize tissue for 3 min at 25Hz (make sure there are no remaining clumps, maybe take longer for muscle).
#Incubate 5 minutes at room temperature.
#Transfer sample to a new 1.5 vial. The ball bearings are reusable after cleaning so do not discard them.
#Add 200 uL Chloroform and shake vigourously by hand for 15s. Do not vortex.
#Incubate at room temperature for 2-3 minutes.
#Centrifuge in a cold eppendorf centrifuge (4C) on maximum for 15 minutes. Sample will separate into a lower, red chloroform phase, an interphase and a colorless upper phase which contains the RNA.
#Add 400 uL of 70% ethanol to a fresh tube.
#Carefully transfer 400 uL of the upper phase to the ethanol tube and mix by vortexing. The remaining chloroform in the previous vial should be disposed of in the phenol waste container under the hood.
#Remove 700 uL of mixture (invert briefly to mix before taking sample) and add to a spin column in a collection tube.
#Spin 15s on max. Discard flow through. Add remaining sample, respin and discard flow through.
#Add 700 uL Wash Buffer I to spin column.
#Spin 15s on max. Discard flow through the collection tube.
#Add 500 uL Wash Buffer II (make sure ethanol was added to the wash buffer) to the spin cartridge.
#Spin 15s on max. Discard the flow through and replace the collection tube.
#Repeat wash by adding 500 uL Wash Buffer II to the spin cartridge.
#Spin 15s on max. Discard the flow through and keep the same collection tube.
#Spin 1 min on max to dry the cartridge. Discard the collection tube and place into a clean recovery tube. Add 100 uL RNAase free water to the center of each tube. This can be adjusted to between 30 and 300 uL elution buffer if necessary.
#Incubate at room temperature for 1min.
#Spin 2 min at 12500 rpm to get purified RNA.
#Quantify the RNA using the nanodrop.

[[Category:qPCR]]
[[Category:Expression]]
[[Category:Transcription]]
[[Category:RNA]]
[[Category:Mouse Work]]
[[Category:Tissues]]

Preparation of RNA Samples from Mouse Tissues

2017-12-14T20:12:23Z

Lgunder:

==Safety Information==
* [[SOP-_Phenol|SOP - Phenol]]
* [[SOP_-_Chloroform|SOP - Chloroform]]

==Materials==
*PureLink RNA Mini Kit (Invitrogen cat#12183-018A)
*Mouse Tissue (50-100 mg, about a 3mm cube)
*TRIZol (Invitrogen cat# 12183-555)
*Chloroform (in solvent cabinet)
*Label tubes, for each sample need 3 eppies, at least one of which is 2 mL and one of which is a recovery tube from the PureLink kit.
*70% Ethanol make with RNAase free water and 100% Ethanol.

==Protocol==
#Add 1 mL TRIzol reagent to each 2 mL tube.
#Cut ~50-100 mg of tissue. If tissue is in RNAlater, cut at room temperature. If tissue is frozen cut on dry ice. Weigh into a fresh 2mL eppendorf tube and keep on ice.
#Using tissue grinder, and after adding the ball bearings to every vial, homogenize tissue for 3 min at 25Hz (make sure there are no remaining clumps, maybe take longer for muscle).
#Incubate 5 minutes at room temperature.
#Transfer sample to a new 1.5 vial. The ball bearings are reusable after cleaning so do not discard them.
#Add 200 uL Chloroform and shake vigourously by hand for 15s. Do not vortex.
#Incubate at room temperature for 2-3 minutes.
#Centrifuge in a cold eppendorf centrifuge (4C) on maximum for 15 minutes. Sample will separate into a lower, red chloroform phase, an interphase and a colorless upper phase which contains the RNA.
#Add 400 uL of 70% ethanol to a fresh tube.
#Transfer 400 uL of the upper phase to the ethanol tube and mix by vortexing. The remaining chloroform in the previous vial should be disposed of in the phenol waste container under the hood.
#Remove 700 uL of mixture (invert briefly to mix before taking sample) and add to a spin column in a collection tube.
#Spin 15s on max. Discard flow through. Add remaining sample, respin and discard flow through.
#Add 700 uL Wash Buffer I to spin column.
#Spin 15s on max. Discard flow through the collection tube.
#Add 500 uL Wash Buffer II (make sure ethanol was added to the wash buffer) to the spin cartridge.
#Spin 15s on max. Discard the flow through and replace the collection tube.
#Repeat wash by adding 500 uL Wash Buffer II to the spin cartridge.
#Spin 15s on max. Discard the flow through and keep the same collection tube.
#Spin 1 min on max to dry the cartridge. Discard the collection tube and place into a clean recovery tube. Add 100 uL RNAase free water to the center of each tube. This can be adjusted to between 30 and 300 uL elution buffer if necessary.
#Incubate at room temperature for 1min.
#Spin 2 min at 12500 rpm to get purified RNA.
#Quantify the RNA using the nanodrop.

[[Category:qPCR]]
[[Category:Expression]]
[[Category:Transcription]]
[[Category:RNA]]
[[Category:Mouse Work]]
[[Category:Tissues]]

Preparation of RNA Sample from Cell Culture

2017-10-25T17:07:28Z

Lgunder: /* Protocol */

==Materials==
*PureLink RNA Mini Kit (Invitrogen cat#12183-018A)
*Mouse Tissue (50-100 mg, about a 3mm cube)
*TRIZol (Invitrogen cat# 12183-555)
*Chloroform (in solvent cabinet)
*Label tubes, for each sample need 3 eppies, at least one of which is 2 mL and one of which is a recovery tube from the PureLink kit.
*70% Ethanol make with RNAase free water and 100% Ethanol
==Protocol==
#Add 1 mL TRIzol reagent to each well of 6 well plate
#Scrape the cells into the TRIzol using an upside down pipet tip
#Pipette the TRIzol plus cells into a 1.5 vial.
#Add 200 uL Chloroform and shake vigourously by hand for 15s. Do not vortex.
#Incubate at room temperature for 2-3 minutes.
#Centrifuge in a cold eppendorf centrifuge (4C) on maximum for 15 minutes. Sample will separate into a lower, red chloroform phase, an interphase and a colorless upper phase which contains the RNA.
#Add 400 uL of 70% ethanol to a fresh tube.
#Transfer 400 uL of the upper phase to the ethanol tube and mix by vortexing. The remaining chloroform in the previous vial should be disposed of in the phenol waste container under the hood.
#Remove 700 uL of mixture (invert briefly to mix before taking sample) and add to a spin column in a collection tube.
#Spin 15s on max. Discard flow through. Add remaining sample, respin and discard flow through.
#Add 700 uL Wash Buffer I to spin column.
#Spin 15s on max. Discard flow through the collection tube.
#Add 500 uL Wash Buffer II (make sure ethanol was added to the wash buffer) to the spin cartridge.
#Spin 15s on max. Discard the flow through and replace the collection tube.
#Repeat wash by adding 500 uL Wash Buffer II to the spin cartridge.
#Spin 15s on max. Discard the flow through and keep the same collection tube.
#Spin 1 min on max to dry the cartridge. Discard the collection tube and place into a clean recovery tube. Add 100 uL RNAase free water to the center of each tube. This can be adjusted to between 30 and 300 uL elution buffer if necessary.
#Incubate at room temperature for 1min.
#Spin 2 min at 12500 rpm to get purified RNA.
#Quantify the RNA using the nanodrop.

Preparation of RNA Sample from Cell Culture

2017-10-25T17:07:01Z

Lgunder: Created page with "==Materials== *PureLink RNA Mini Kit (Invitrogen cat#12183-018A) *Mouse Tissue (50-100 mg, about a 3mm cube) *TRIZol (Invitrogen cat# 12183-555) *Chloroform (in solvent cabine..."

==Materials==
*PureLink RNA Mini Kit (Invitrogen cat#12183-018A)
*Mouse Tissue (50-100 mg, about a 3mm cube)
*TRIZol (Invitrogen cat# 12183-555)
*Chloroform (in solvent cabinet)
*Label tubes, for each sample need 3 eppies, at least one of which is 2 mL and one of which is a recovery tube from the PureLink kit.
*70% Ethanol make with RNAase free water and 100% Ethanol
==Protocol==
#Add 1 mL TRIzol reagent to each well of 6 well plate
#Scrape the cells into the TRIzol using an upside down pipet tip
#Pipette the TRIzol plus cells into a 1.5 vial.
#Add 200 uL Chloroform and shake vigourously by hand for 15s. Do not vortex.
#Incubate at room temperature for 2-3 minutes.
#Centrifuge in a cold eppendorf centrifuge (4C) on maximum for 15 minutes. Sample will separate into a lower, red chloroform phase, an interphase and a colorless upper phase which contains the RNA.
Add 400 uL of 70% ethanol to a fresh tube.
#Transfer 400 uL of the upper phase to the ethanol tube and mix by vortexing. The remaining chloroform in the previous vial should be disposed of in the phenol waste container under the hood.
#Remove 700 uL of mixture (invert briefly to mix before taking sample) and add to a spin column in a collection tube.
#Spin 15s on max. Discard flow through. Add remaining sample, respin and discard flow through.
#Add 700 uL Wash Buffer I to spin column.
#Spin 15s on max. Discard flow through the collection tube.
#Add 500 uL Wash Buffer II (make sure ethanol was added to the wash buffer) to the spin cartridge.
#Spin 15s on max. Discard the flow through and replace the collection tube.
#Repeat wash by adding 500 uL Wash Buffer II to the spin cartridge.
#Spin 15s on max. Discard the flow through and keep the same collection tube.
#Spin 1 min on max to dry the cartridge. Discard the collection tube and place into a clean recovery tube. Add 100 uL RNAase free water to the center of each tube. This can be adjusted to between 30 and 300 uL elution buffer if necessary.
#Incubate at room temperature for 1min.
#Spin 2 min at 12500 rpm to get purified RNA.
#Quantify the RNA using the nanodrop.

Thawing Culture Cells

2017-09-27T14:36:46Z

Lgunder:

Materials
* Media( High Glucose DMEM: Pre-prepared 500mL: 1% PSG and 10% PBS)
*C2C12 cells
Protocol
* Thaw vile of C2C12 cells in 37C water bath approx. 1 min
* Pipette 10mL of media to plate in sterile hood
* Pipette cells from vile into plate with media (evenly pipette/distribute throughout plate)
* Lightly swirl the plate until bottom of plate is covered
* Review cells under microscope
* Incubate plate in 37C incubator
* Check cells in 4-6 hours for growth (under microscope)
* Replace media after significant growth approx. 85-90%
Notes:
Sanitize surface, vials, hood etc with 70% ethanol and paper towel,
Always return plates to incubator,
Trash all cell vials and waste in contact into biohazard trash for autoclave,
Review cells under microscope for growth periodically
See Relevant links
[[Splitting Cells]]
[[Culturing and Differentiating C2C12 Cells]]

Thawing Culture Cells

2017-09-27T14:30:41Z

Lgunder: Created page with "Materials * Media( High Glucose DMEM: Pre-prepared 500mL: 1% PSG and 10% PBS) *C2C12 cells Protocol * Thaw vile of C2C12 cells in 37C water bath approx. 1 min * Sterilize vile..."

Materials
* Media( High Glucose DMEM: Pre-prepared 500mL: 1% PSG and 10% PBS)
*C2C12 cells
Protocol
* Thaw vile of C2C12 cells in 37C water bath approx. 1 min
* Sterilize vile and media bottle with 70%ethanol
* Add 10mL of media to plate in sterile hood
* Pipette cells from vile into plate with media (evenly pipette/distribute throughout plate)
* Review cells under microscope
* Incubate plate in 37C incubator
* Check cells in 4-6 hours for growth (under microscope)
* Replace media after significant growth approx. 85-90%
Notes:
Sanitize surface, vials, hood etc with 70% ethanol and paper towel,
Always return plates to incubator,
Trash all cell vials and waste in contact into biohazard trash for autoclave,
Review cells under microscope for growth periodically
See Relevant links
[[Splitting Cells]]
[[Culturing and Differentiating C2C12 Cells]]