Protocols

Cutting Bands from a Gel

Cutting the Bands

  • Fill appropriate number of 500 µL PCR tubes with 50 µL of 10 mM Tris buffer
  • Carefully slide the gel onto the wetted transilluminator plate (wetting the edges of the gel is helpful)
  • Wearing protective equipment, turn on the transmitted UV switch
  • Using a sterilized scalpel (cleaned with 70% ethanol), cut along the top and bottom edges of the band
  • Snip the left and right edges of the band with sterilized forceps (cleaned with 70% ethanol)
  • Remove the excised band and place in corresponding PCR tube
  • Repeat as necessary
  • Store tubes at 4oC for three to five days to allow DNA to diffuse into the buffer

DGGE Protocol

Preparation of Acrylamide Solutions

0% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Milli-Q Water 124.5mL
    • Combine first two components
    • Bring volume up to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

80% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Formamide (freshly deionized) 48mL
Urea 50.4g
Milli-Q Water ~75mL
    • Combine first four components (urea dissolves slowly)
    • Bring volume to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

Deionization of Formamide

Formamide 70mL
Mixed Bead Resin 1-2 spoonfulls
  • NOTE: Formamide must be kept wrapped in foil throughout its use due to its sensitivity to light
  • Combine formamide and resin
  • Mix on stir plate for 60 minutes
  • Use Whatman filter paper to remove resin from formamide

Cleaning the Glass Plates

  • Remove one set of glass plates (on tall and one short) and two spacers from the soap solution
  • Wash both sides thoroughly with dish soap and rinse with tap water followed by distillied water
  • Wash the gel-side of each plate with the following reagents in specified order:
    • Windex
    • Acetone
    • 70% Ethanol

Plate Assembly

  • Place the spacers along the edges of the tall glass plate (gel-side up), with the notches facing towards the center of the plate
  • Place the short glass plate (gel-side down) onto the spacers. Align the assemblage with all sides flush, making sure spacers extend slightly past the glass plate edges
  • Lightly tighten the supports to either side of the assemblage, ensuring that the arrows on the supports are facing up and towards each other
  • Place the glass plate assemblage onto the positioning area of the already leveled base
  • Release the tension of the supports and position the plates and the spacers so that they are flush against each other. Proper alignment of the plates is essential for the formation of a good seal
  • Tighten the supports to lock the assemblage into place
  • Pull out the knobs on either side of the base and insert the plates into the pouring area, ensuring that the assemblage is flush against the grey strip
  • Push in the knobs and rotate the screws in a counterclockwise direction (as looking straight down) to secure the plates
  • Verify that there is some bowing in the grey strip to ensure a proper seal

Pouring the Gel

  • Make three solutions from the following chart in 15 mL centrifuge tubes (one of these must be the 0% plug)
0% 10% 20% 30% 40% 50% 60% 80%
 0% solution 6 mL 9.625 mL 8.25 mL 6.875 mL 5.5 mL 4.125 mL 2.75 mL 0 mL
80% solution 0 mL 1.375 mL 2.75 mL 4.125 mL 5.5 mL 6.875 mL 8.25 mL 11 mL
APS (10%) 44 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL
TEMED 6.0 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL
  • Set up the puring apparatus, making sure that the valve connecting the two wells of the gradient maker is closed (~30o from horizontal)
  • Turn on stir bar to 6
  • Center the comb in between the plates and place the syringe between the plates and against one of the spacers
  • Thoroughly mix the acrylamide solutions (20% with either 40% or 60% or 80%) with the APS and TEMED
  • As quickly as possible, pour the higher concentration solution into the front chamber (the one closest to the outlet valve) and pour the lower concentration solution into the back chamber
  • Open the valve between the wells on the gradient chamber (bring to horizontal position) and begin the mixing and the pouring of the gel
  • To promote even mixing, reduce the speed of the stir bar as the gel pours (one should just barely be able to see the mixing in the front chamber). If the stirring speed is too great, the higher concentration denaturant will flow back into the other chamber
  • When the wells are approximately 75% empty, thoroughly combine APS, TEMED and the 0% solution. Add this mixture to the rear well as the front one empties
  • Once both wells are empty, clean the pouring apparatus by running Milli-Q water through the system multiple times
  • Allow the gel to sit for at least two hours. If leaving overnight (recommended), cover assemblage with a large ziploc bag and store in a large tupperware container with a beaker full of water (to reduce dessication)

Assembling the D-Code (Gel Running) System

  • Clean off a set of glass plates (dish soap only)
  • Line the space between the plates with Parafilm
  • Lock plates into place with supports, ensuring they are flush
  • Carefully remove the comb from the gel, along with any excess gel from around the wells
  • With the tall glass facing out and short glass facing in, snap both assemblages into the holder
  • Fill D-Code system with approximately 7 L of 1xTAE (140 mL 50xTAE and 6860 mL Milli-Q water) until fill-line is reached
  • Place the holder down into the buffer, ensuring that it is properly in place
  • Place the top of the D-Code system on top of the apparatus. Verify that all electrical connections are created
  • Plug in electrical chord and turn power switch on
  • Set temperature to 65oC and turn on the heater and pump. Make sure that the buffer level rises and remains above the negative electrode wire
  • Allow D-Code to reach 65oC (90 to 120 minutes)

Loading the Gel

  • Turn off power and remove top of D-Code system
  • Prepare samples by combining equal amounts of template (at least 300 ng) with 5x loading buffer in 500 µL PCR tubes
  • Using a P-100 and corresponding gel loading tips, load samples by placing the pipettor tip half-way into the well and expunging sample with extreme care (loading the wells is made easier by avoiding air bubbles at the pipette tip and expunging slowly)
  • Load the standard into every fourth lane
  • Replace top to the D-Cody system, ensuring all electrical connections are made
  • Plug in D-Code and plug it into both the electrical outlet and the BioRad PowerPack300 (with DCODE chip)
  • Turn on D-Code system and set temperature to 60oC
  • Turn on heater
  • Set PowerPack300 (with DCODE chip) to 200V
  • Turn on the pump only after samples are pulled into the gel (approximately 15 minutes)
  • Let gel run for 3.5 to 4 hours at 200V (or 12 hours at 100V)

Staining and Imaging the Gel

  • Drain the excess buffer from the top of the gel and dismantle the apparatus
  • With the upmost of caution, remove the supports and spacers
  • With extreme care, use one of the spacers as a lever to pry off the short glass plate, leaving the gel on the tall glass plate
  • Soak the tall plate (with gel) in an ethidium bromide stain, and place on rocker (slow mode) for 30 minutes
  • Destain in Milli-Q water for 15-20 minutes
  • Thoroughly wet the imager’s glass plate with water
  • Tilt the glass plate onto the wet surface of the imager, allowing the gel to slowly slide off (continually wetting the edges of the gel is helpful)
  • Once imaging completed and bands to be cut are identified, remove the gel by re-wetting the glass plate and carefully sliding the gel onto it
  • Transport gel to transilluminator for band cutting

DGGE Standards

Materials

ND1-1
ATTC 29133
S1006 (diatom)
MoBio Plant DNA Extraction Kit

or

Materials for PCI Method

Procedure

Extract DNA of all samples using either the MoBio Plant Kit or PCI Extraction method

Amplify DNA with cyanobacterial GC-clamped primers using the standard 30 cycle amplification for cyanobacteria

NOTE: These strains have all been shown to give only one band

Hazardous Waste Pickup

Make sure waste is appropriate, as Risk Management will only accept:

  • Chemicals
  • Biowaste
  • Lamps
  • Batteries

Make sure container is appropriate

  • Condense all waste into as few containers as possible
  • All labels on bottles must be crossed out
  • Bottles must have tight, secure lids
  • DO NOT use biohazard bags for any waste that is not comprised of human tissue or human pathogens
  • Verify that bags are free from tears and leaks

Get a Hazardous Waste Tag (two drawers below hotplate in LSE422)

Fill tag out completely and legibly or waste will not be picked up

  • Chemical: list all chemicals in container using written out names (not chemical formulas)
  • ~~NOTE- they will not pick up any unknowns; everything must have a name~~
  • Amount: rough approximations are fine
  • Vol%: if known
  • Generator: Ferran Garia-Pichel
  • Phone: 727-7762
  • Date: date tag filled out
  • Dept: Microbiology
  • Building: LSE
  • Room: 404, 422, or 427
  • Category:
  • Solid/Liquid: this should reflect container’s contents, not actual contaminant
  • pH: if known

Attach tag to container and set in a visible area where it will not be moved

Contact Risk Management via

Be prepared to provide the following information

  • Your name
  • Your phone number
  • Location of the waste (building, room number, and location within that room)
  • Description of the waste
  • Description of the containers (both type and number)
  • Confirmation that the waste has been tagged
  • Any questions regarding either the waste or the filling out of the tag

Expect to wait a few days before waste is picked up. If nobody comes for it, or it needs to get picked up quickly, call daily

Imaging and Quantifying Gels

Staining

  • Inspect stain and destain solutions to make sure that they are fresh and free of broken gel pieces
  • To make new stain :
    • Pour out existing stain into ethidium bromide waste container
    • Rinse plastic with MilliQ water, pouring rinse into waste container
    • Fill plastic with approximately one liter of MilliQ, and add a few drops of ethidium bromide until water is slightly colored
  • To make new destain:
    • Pour out existing destain into ethidium waste container
    • Rinse plastic with MilliQ water, pouring rinse into waste container
    • Fill plastic with appoximately one liter of MilliQ water
  • Soak the gel in ethidium bromide stain for 30 minutes
  • Use spatula to move gel into a Milli-Q water destain, and let sit for 15 minutes

Positioning Gel

  • Turn on FluorS-MultiImager (power switch is located on back-right hand side)
  • Thoroughly wet the imager’s glass plate with water
  • Remove gel from destain, rinse with Milli-Q water to remove any loose pieces of agarose, and move onto the wet surface of the imager
  • Change gloves
  • Wearing clean gloves, sit at computer and click on “Quantity One” icon (a red icon, found most often on the upper right side of the Desktop)
  • Select “fluorese” from “File” menu
  • Click on “Position” button
  • Use the red plus-mark and the blue grid to center and align the gel
  • Click on “Stop” button and close door to FuorS-MultiImager

Creating Image

  • Set exposure time to 10.0 and click on “Preview” button
    • If bands are too faint, increase exposure time and re-preview
    • If gel is too bright, either decrease exposure time or destain longer
  • Click on “Acquire” button
  • Save image by selecting “Save” from “File” menu, and/or print image by going to “Print” (also under the “File” menu) and selecting “Print Image,” then clicking on “Go” button

Quantifying Bands

  • Open “Lane Tools” and “Band Tools” toolbars (16th and 17th icons from left, respectively, on main Quantity One toolbar)
  • Zoom in on gel (7th icon from left, on main toolbar) as much as possible without cutting off any bands
  • Mark all lanes of interest
    • Select “Create Lane” (6th icon from left, on Lane Tools)
    • Position the mouse above the bands in first lane of interest, centering it horizontally
    • Click, hold, and drag the mouse down past the end of the last band within that lane
    • There should now be a red line running through the center of each band within the first lane
    • Click on “Adjust Lane” (7th icon from left, on Lane Tools)
    • Position mouse over least centered section of line
    • Click, hold and drag the mouse to adjust the line as needed
    • Repeat with all relevant lanes
  • Click on “Detect Bands” (1st icon from left, on “Band Tools”), click on “Close” (all bands should now have brackets roughly surrounding them)
  • Click on “Remove Band” (4th icon from left, on Band Tools)
  • Click on a set of brackets that does not correspond to an actual band, click on “Yes,” and repeat for all other incorrect brackets
  • Adjust brackets vertically
    • Click on “Adjust Bands” (3rd icon from left, on “Band Tools”)
    • Click and hold on the upper bracket corresponding to the first band
    • Adjust bracket both visually and by using the pop-up graphs
    • VISUALLY: The bracket should serve as the upper/lower limits to the bands
    • GRAPHS: Each band has its own peak on the graph. The brackets should be set so that only the steepest section of each peak is highlighted in yellow.
    • NOTE: In theory, each one of these approaches should yield the same result. In practice, such is rarely the case, so it is important to rely on a combination of the two.
  • Adjust brackets horizontally
    • Click on “Lane Width” (10th icon from left, on “Lane Tools”)
    • Click on first lane
    • Arbitrarily adjust number and click “OK”
    • Continue this process until brackets serve as borders to the bands
    • If the (red line demarking the) lane is not centered over the bands it will have to be re-aligned, as it will be impossible to properly adjust the brackets otherwise.
    • NOTE: Re-aligning the lanes will erase the brackets. This time, when preparing to “Detect Bands,” change the “Lanes” option from “All” to “One”, and type in the re-adjusted lane’s number, as labeled by Quantity One.
  • Select “Quantity Standards” from “Analysis” menu, and click on “Create New”
  • Click on the button corresponding to “Select Bands”
  • Return to the window with the gel imange and click on bands from lane containing the mass standard
  • Return to the “Quantity Standards” window
  • Set “Interpolation” to “Linear Regression,” and “Extrapolation” to “Yes”
  • Manually alter the “Quantity” column of the table at the bottom of the window to reflect the known values (if four lanes appear, the values are 20, 15, 10, and 6 for each 1µL added)
  • Click on “Show Curve”
  • If any point is very off, remove it from the chart
  • Click on the button corresponding to “Calibrate Gel,” and click on “OK”
  • Select “All Lanes Report” from “Report” menu, and select “Report”
  • Print report if necessary by choosing “Print All” (2nd icon from left, at bottom of window)
  • Save calculations in personal folder by selecting “Save” from “File” menu (calculations and gel image are all saved together)
  • NOTE: Quantity One caculates the concentration (see “Calibrated Quanitity” column in the “Report” printout) in terms of ng/mL
  • Quit program

Cleanup

  • Remove gel from FuorS-MultiImager and either store in Ziplock bag or throw away in ethidium bromide waste container
  • Clean off FluorS-MultiImager with ethanol
  • Remove any broken pieces of gel from stain and destain, place in ethidium bromide waste container
  • Throw gloves away in ethidium bromide waste container

Ordering Information

If you are not the Research Technician

  • Decide what you want
  • Get a catalogue from outside LSE422 and look up pertinent information :
    • Actual and complete item name,
    • Company name, mailing address, and phone number,
    • Catalogue number,
    • Unit sold (eg each, packet of six),
    • Price per unit
  • Neatly put the catalogue back in alphabetical order (ie where you got it)
  • Give this information, along with amount desired to the technician along with any special concerns (eg rush delivery)
  • Wait 7 to 10 days

NOTE: you must provide the aforementioned information to the technician even if you place an order with the credit card

If you are the (acting) Research Technician

  • Make sure you are provided all necessary infomation from the grad student/professor
  • Get the order approved by Dr. Garcia-Pichel, and ask which grant to use for the order
  • Get a green departamental order form from the file cabinet
  • Fill out the form, using completed ones as templates to answer any questions
  • Get Dr. Garica-Pichel to sign it
  • Photocopy the form, and turn the original over to Kristy Bolen (LSE 210)
  • File the photocopy according to grant number under the “incomplete” heading
  • Upon arrival, double check accuracy and condition of item(s)
  • If there are no complications:
    • Write “received okay” on packing slip along with date
    • Photocopy slip
    • Give original slip to K. Bolen
    • Attach photocopy of slip to photocopy of order form
    • File photocopies appropriately (according to year and account number)
  • If there are complications:
    • Leave order as inctact as possible
    • Find order form and double check order
    • Take both packing slip and order form to K. Bolen (LSE 210) and explain the problem
    • From here, everything will be situation-specific
  • It is your responsibility to keep track of orders and make sure they arrive. Many companies will not contact ASU if there is a problem with the order, be it their fault or ours.

PCI Extraction

Reagents and Solutions

TESC Buffer 11 mL/sample
Proteinase K 30 µL/sample
10% SDS 550 µL/sample
PCI ~15 mL/sample
CI ~10 mL/sample
Isopropanol ~6 mL/sample
70% Ethanol (-20oC) ~0.5 mL/sample
10 mM Tris or TE Buffer 250 µL/sample

 

Materials

Scalpel, autoclaved 1
Tweezers, autoclaved 1
Weigh Boats, autoclaved 1 per sample
All-Glass Tissue Grinder or ceramic mortar and pestle, autoclaved 1 per sample
1.5 mL Eppendorf Tubes, autoclaved 1 per sample
100 µL Pipette 1
100 µL Pipette Tips, autoclaved one per sample
1 mL Pipette 1
1 mL Pipette Tips, autoclaved many
5 mL PCI-Pipette (located in the hood in LSE404) 1
5 mL Pipette Tips, autoclaved many
Water Baths one a 37oC
one at 50oC
one at 65oC
50 mL Plastic Centrifuge Tubes with Screw Caps four per sample

Sample Preparation

  • Autoclave everything
  • Using sterile tweezers, scalpel and weigh boat, weigh out approximately 50-300 mg wet mass of sample (flame sterilize tweezers and scalpel between samples)
  • Transfer sample into all-glass tissue grinder (potter), add 1 mL TESC buffer and homogenize
  • Using 1 mL pipette, transfer homogenate to 50 mL centrifuge tube
  • Rinse potter with 4 mL TESC buffer and combine with above homogenate
  • Rinse potter with 1 mL TESC buffer and combine with above homogenate
  • Freeze at -80oC (~20 minutes), thaw at room temperature for ~5 minutes, thaw completely in 65oC water bath (~10 minutes)
  • Repeat above sequence five times (for a total of six freeze-thaws)

Extraction

  • Add 30 µL Proteinase K solution and 550 µL 10% SDS solution to sample
  • Gently mix by inverting capped tube several times
  • Incubate at 50oC for 40 minutes, gently mixing every 10 minutes
  • Add 5 mL PCGently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 10 minutes
  • Carefully transfer aqueous (ie top) layer into a new 50 mL centrifuge tube
  • Add 5 mL TESC buffer to original tube
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 10 mintues
  • Combine aqueous (ie top) layer with that from above
  • Add one volume (~10mL) PCI to combined aqueous layers
  • Gently mix by inverting capped tube several times
  • Incubate at 65oC for 10 minutes, gently mixing every five minutes
  • Centrifuge at 4,100 rpm for 12 minutes
  • Transfer aqueous (ie top) layer into a new 50 mL centrifuge tube
  • Add one volume (~10mL) CI
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 12 minutes
  • Transfer aqueous layer to new 50 mL centrifuge tube
  • Centrifuge at 4,100 rpm for 12 minutes
  • Remove any remaining PCI/CI from tube
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,000 rpm for 8 minutes

Culmination

  • Precipitate nucleic acids with 0.6 volume (~6 mL) isopropanol for at least 30 minutes at room temperature, or overnight at 4oC
  • Alternatively, add 0.04 volume (~400 µL) 5 M NaCl and mix, add 2 volumes (~20 mL) cold (-20oC) 100% ethanol and mix, precipitate for 60 minutes in an ice bath
  • Centrifuge at 4,100 rpm for 60 minutes at 4oC
  • Decant supernatant
  • Wash pellet with cold (-20oC) 70% ethanol, pouring it slowly along the tube’s wall. If pellet dislodges, re-centrifuge at 4,100 rpm for 20 minutes at 4oC
  • Dry pellet for 1-3 hours in vacuum
  • Add 250µL 10 mM Tris (or TE) buffer to tube
  • Incubate at 37oC for at least 30 minutes to allow for dissolution
  • Transfer re-suspended pellet into a 1.5 mL Eppendorf tube
  • Store at -20oC

PCR

Materials

  • PCR tubes (one per sample, two for controls, and one extra)
  • Two 2 mL microcentrifuge tubes
  • Template
  • E. coli DNA (or other positive control)
  • Primers, forward and reverse
  • 10% BSA (LF)
  • 10x Ex Taq buffer
  • dNTP mixture
  • 5x Eppendorf Enhancer
  • Milli-Q water, autoclaved
  • Ex Taq DNA polymerase
  • Assorted pipettors and tips

Procedure

  • Open all PCR and microcentrifuge tubes and place in racks found in the PCR hood, along with all pipettors and tips (note: all pipettors should already be in hood)
  • Close the side panels and set the UV light timer for 20 minutes
  • Turn on UV light and blower for the flow hood
  • Gather all reagents and samples, placing them in an ice bath
  • Enhancer should be placed in a warm water bath to dissolve precipitate
  • Enter relevant data into the “Master Mix” calculation spreadsheet; make a print out
  • In PCR hood, prepare master mix in one 2 mL microcentrifuge tube
  • Aliquot master mix into PCR tubes
  • Transfer PCR tubes to ice bath and move to flow hood
  • Add template to each reaction tube, E. coli (or equivalent) to positive control, and PCR water to negative control
  • Place caped and labeled tubes in PCR thermocycler, then close lid
  • Turn on thermocycler, select desired PCR cycle, and begin
  • Pause thermocycler at very end of second cycle
  • Quickly add Taq polymerase to each tube (note: Do not expel polymerase from tip; it will flow out automatically. Do not allow any air bubbles to form in PCR tube.)
  • Close thermocycler lid and press “resume”
  • Store tubes at 4oC

PCR Cycles

Cyanobacterial Primers (cyano)

First Cycle (run once)

  • 95oC for 5 minutes
  • 80oC for 1 minute
  • With about 3 seconds remaining, pause thermocycler and add Taq

Second Cycle (run 30 times)

  • 94oC for 1 minute
  • 60oC for 1 minute
  • 72oC for 1 minute

Third Cycle (run once)

  • 72oC for 9 minutes
  • 4oC hold

Eubacterial Primers (eubac td)

First Cycle (run once)

  • 95oC for 5 minutes
  • 80oC for 1 minute
  • With about 3 seconds remaining, pause thermocycler and add Taq
  • 65oC for 1 minute
  • 75oC for 3 minutes

Second Cycle, Touchdown PCR (run 20 times)

  • 94oC for 1 minute
  • 65oC for 1 minute
  • 75oC for 3 minute
  • NOTE: this is a touchdown cycle; the temperature for the second step must be lowered by 1oC every second cycle (ending on 55oC).

Third Cycle (run 9 times)

  • 94oC for 1 minute
  • 55oC for 1 minute
  • 72oC for 3 minute

Fourth Cycle (run once)

  • 72oC for 7 minutes
  • 4oC hold

Converting Sequencing Files

Converting from PC to Mac

  • Save/transfer all PC files into a single folder on the computer
  • Open ConvProg (desktop alias named “Sequence Converter”)
  • A standard folder select dialog box will appear
  • Navigate to the folder containing all files to be converted (note: do not open said folder, simply select it)
  • Click the “choose” button

Agarose Gel Protocol

Assembling Electrophoresis Cell

  • Gather all parts. These should be located on the shelf above the pipette rack, but can often be found in/near the sinks or in the 4oC : Base, Gel-Casting Gates (two), Gel Tray,Comb(s),Safety Lid with Electrical Cables
  • Place the base on an open part of the counter
  • If base is not level, adjust Leveling Feet accordingly
  • Insert the Gel-Casting Gates into their corresponding grooves
  • Place the Gel Tray between the Gel-Casting Gates
  • NOTE: there is a nice diagram in the BioRad Life Science Research Products catalog on page 255

Preparing the Gel

  • Combine either 40 mL buffer with 400 mg agarose in an Erlenmeyer flask for a small gel or at least 100 mL buffer with 1.0 g agarose, to create a 1% solution
  • Use 1x TAE as your buffer if your samples are DNA extracts, and 1x TBE if your samples are PCR amplified DNA
  • Plug up flask with a paper towel
  • Microwave flask for roughly one to two minutes for a small gel and two to three minutes for a large gel to dissolve agarose, stopping microwave and swirling flask every time it shows signs of boiling
  • Once completely dissolved, cool solution under running water until the flask can be held without burning skin
  • Double check that Gel-Casting Gates are securely in place and pour agarose solution onto Gel Tray
  • For small gels, pour all 40 mL; for large gels, fill between two and three ticks (these are on the sides of the base)
  • Rinse out Erlenmeyer flask, fill with water, and set aside
  • Pop any bubbles in gel with a syringe needle
  • Insert appropriate Comb(s) into the appropriate Comb Slots
  • Allow gel to solidify
  • Thoroughly clean out Erlenmeyer flask
  • Gel can now be saved for later use. Place in Ziplock bag and store at 4oC

Loading Samples

  • Remove Comb(s) and Gel-Casting Gates
  • Wipe gel off Comb(s) and Gel-Casting Gates with paper towel to remove any agarose residue
  • Fill base with same buffer as above to cover gel
  • On a strip of Parafilm, measure out 5 µL sample and 2 µL Nucleic Acid Sample Loading Buffer, 5x
  • Mix sample and loading buffer by slowly sucking in and out of pipettor tip
  • Load samples into wells
  • Never use outside lanes
  • Dip pipette tip into buffer on side with red Bananna Plug/Electrode Cassette before loading well to remove any excess dye
  • Carefully lower tip half-way into well, being careful not to touch gel
  • Load well slowly to ensuring none of the sample escapes into the buffer
  • Raise pipette tip out of buffer before releasing thumb (very important)
  • Load 5 µL Precision Molecular Mass Standard into one well and 1 µL into another

Running Gel

  • Place Safety Lid on Base, matching up Electrical Cables to Bananna Plugs/Electrode Cassettes according to color
  • Plug Electrical Cables into power source (PowerPack 300 or EC105)
  • Turn on power source, set voltage to 110V for small gels and 120V for large gels, and press the “running man” button to begin (verify that the gel is running by looking for a stream of bubbles originating at the black Bananna Plug/Electrode Cassette)
  • Run small gels for approximately 20 minutes and large ones until the dye is 75% across the gel
  • Turn off and unplug power source, and disconnect Electrical Cables
  • Remove Safety Lid and Comb(s)
  • Remove Gel Tray with gel on it (gel can either be imaged or stored in a Ziplock bag at 4oC)
  • Pour buffer out of Base into a labeled container and store at 4oC for later use
  • Clean out Base, Comb(s), and Gel Tray

Imaging Gel and Quantifying Bands

  • Follow standard procedure