Sterile Technique Video on Google Drive

Overview

    Sterile Technique is an essential aspect of successful cell culture practice. When growing and passaging cells in a lab, the cells are at a high risk for becoming contaminated with bacteria, fungi, or other contaminants if sterile technique is not followed. If a cell line becomes contaminated or infected, the cells will likely die soon after and your other cell lines are at risk of also becoming contaminated. For this reason, sterile technique must be deliberately followed for every task that is performed handling cells. This includes, but is not limited to, passaging cells, plating cells for transfection, transfecting cells, oberving cells under a microscope, and any other protocol that requires the physical handling of cells in a Petri dish or well plate.

A general overview of sterile technique can be seen in this Video by Thermo Fisher

• This video obviously is not filmed in our lab, and slight variations are seen in the video when comparing cell culture in our lab. In general, all techniques mentioned in this video should be accounted for when handling cells in our lab. When handling our current cell lines (LM45 and LM45-CD47), it is not required to wear a protective lab coat or lab glasses, but you may do so if you have them at your disposal. Additionally, the pipettes we currently use for cell culture are the same as the pipettes used in our lab for general purposes.

Sterile Technique In Our Lab

    When handling cell lines in the our lab in any fashion, the following rules must be adhered to:

• Always wash hands well with soap prior to handling cells and putting gloves on

• Following hand washing, always wear gloves (purple latex gloves found near door in lab) prior to handling cell culture dishes/any cell culture materials

  • After putting gloves on to clean hands, spray gloves with 70% ethanol and rub gloved hands together until dry

• If Using Cell Culture Hood (Room 504)

  • Prior to using hood, turn on the light to the side labeled "fluorescent" and turn on the blower

    • After this is done, spray down the entire surface of the hood with 70% ethanol, and wipe down the hood in one continuous motion with a paper towel

      • Motion consists of starting in one back corner of the hood, moving horizontally along the back of the hood, then moving back towards the front of the hood in a horizontally snaking pattern

  • Acquire all materials necessary for your cell culture tasks that day prior to grabbing cell culture dishes that already contain cells

  • Spray down/Wipe down any cell culture materials being used with 70% ethanol prior to putting in hood

    • Note: ethanol removes sharpie, so be aware of how tubes are labeled prior to wiping with ethanol and replace labels afterward if needed

  • Always place the lid of any container used in the hood face down (interior of the lid facing down)

  • You may use micropipettes used for general purposes in the lab, but they must be sanitized with ethanol prior to placement in the hood.

  • ONLY USE micropipette tips designated for cell culture. These can be found inside of the cabinets in the back of the lab above the oocyte injecting station or above the centrifuge. These pippete tips have been autoclaved and are sterile. DO NOT OPEN THE PIPETTE TIP BOX OUTSIDE OF A STERILE CELL CULTURE HOOD.

    • If a new box is being used, ensure the tips have been autoclaved prior to their use.

  • Always discard micropipette tips after a single use

  • ****Only ever open micropipette tip boxes, solutions used for cell culture, and any cell culture dish inside of the previously sanitized hood. 

• Cell Incubator (Located in Room 504)

  • The cell incubator door should remain properly sealed at all times, and if it needs to be opened, the door should remain open for only as long as it needs to

  • ONLY CELL CULTURE DISHES/WELLED PLATES ARE TO PLACED IN THIS. NO BACTERIAL PLATES OR ANY OTHER BIOLOGICAL TOOLS/CHEMICALS MAY GO IN THIS INCUBATOR

  • Sanitize the shelves of the cell incubator with Clorox/lysol disinfecting wipes periodically or if contamination of the shelves is assumed

Passaging Cells

Passaging Protocol (Video)

Materials Needed:

    - Plates containing the cells to be passaged

    - Fresh, Sterile cell culture plates to passage the cells into

    - Complete Growth Media

    - Trypsin

    - 1 mL and 200 uL micropipettes and STERILIZED micropipette tips

    - Cell Culture Dishes (35 mm or 60 mm)

    - Sharpie/Tool for Labeling Plates

    - Erlenmeyer flaşk for storing old growth media

Protocol

1. Wash hands and put on gloves, spray gloves with ethanol.

2. Turn on the blower and light for the hood and open it

3. Clean the hood with ethanol, wiping from back to front

4. Procure complete growth media (RPMI 1640 + 10% FBS) and trypsin from fridge and place in hood

5. Place previously autoclaved micropipette tips of all three sizes in hood along with the pipettes

6. Unscrew the caps to the media and trypsin but leave them on 

7. Obtain cells from the incubator and place in the hood 

8. Check viability of each dish of cells by looking under microscope

   1. Cells should be 100% confluent

   2. Refer to Photos of 100% Confluent Melanoma Cells Before Passaging

9. Place new cell dishes in hood and label with name and date

10. Remove the lid of the dish that currently contains cells and place lid facing down, tilt dish at a 45° angle to have the media collect in one spot.

11. Pipette out the media from each dish using the suction pipette/aspirator, made sure to not let the pipet tip touch anything but the plate (not your gloves) and use a new pipet tip for each plate.
    - If no aspirator/suction pipette is available, you may use a sterile 1 mL pipette (with sterile tips) to remove the old growth media. The old growth media can be removed from the plates and put into an Erlenmeyer flask. Once all old growth media is collected, the Erlenmeyer flask can be sealed with aluminum foil and then autoclaved (to deactivate the antibiotics and sterilize the growth media) after passaging is complete. Once autoclaved, the old growth media can be dumped down the drain.

12. Add the appropriate amount of trypsin to each dish 

    1. 400 uL to 60 mm dish 

    2. 200 uL to 35 mm dish

13. Allow the cells to incubate in the trypsin in the hood for 5 minutes

14. During this incubation period, add the appropriate amount of complete growth media to the new dishes 

    1. 4 mL to 60 mm dish

    2. 2 mL to 35 mm dish

15. After incubation period, angle the parent dish at 45o and suction trypsin using a micropipette and wash the cells down the side to diminish any remaining adherence

16. Triturate to mix the cells

Note: triturate means to slowly pipette the cells up and down to mix them into a homogeneous solution, try not  to introduce any air bubbles into the solution

17. Add an appropriate number of drops of triturated trypsinized cells from the parent plate to the new plate.

    1. For 60 mm dishes, use approximately half a drop of a 200 uL pipette

    2. For 35 mm dishes, use 1 drop of a pippete set to 10 uL (black pipette with red plunger)

18. Spread the cells all over the new plate by pipetting up and down

19. Gently move the plate from side to side, up and down, to evenly spread the cells

    1. Do not move the plate in a circular motion or the cells will collect in the middle of the dish and will not be evenly spread out.

20. Check the new plate under a microscope to ensure the cell amount and distribution is sufficient. If there isn’t enough cells, add another half drop/drop of cells from the trypsinized dish, spread out cells, and observe under microscope again until desired cell count in new dish is observed.

21. Discard the old cells in the trash

Pre-transfection plating

The video will give you a mostly accurate description of how to pre-plate for transfection. The written protocol has been updated since Blake's video to include Alec's transfection log and calculator, as well as the use of transfection growth media (using Opti-MEM) instead of complete growth media (using RPMI 1640).

***Note: As with all cell culture procedures, ensure sterile technique is properly followed. This includes wearing gloves with clean hands, sterilizing the hood before using, only opening plates, micropipette tips boxes, and solutions inside of the sterile hood, and only putting cells in sterile plates/tubes.

Materials Needed:

- Plates containing the cells desired for transfection

- Transfection growth media (Opti-MEM with 2.5% FBS)

- Trypsin

- Hemocytometer

- 1 STERILE falcon tube (15 or 50 mL)

- 1 STERILE eppendorf tube (capable of holding at least 1 mL)

- 1 mL, 200 uL, and 10 uL micropipettes and STERILIZED micropipette tips

- 12 well plate

- Sharpie/Tool for labeling wells

Protocol

1. Before beginning, ensure you have marked your transfection plans on the Transfection Log. Include the plasmid, the dose of lipofectamine, and the media solution (usually Opti-MEM) used in each well. This should go on the Log tab and be marked with the date.

2. Roughly 24 hours before transfection, remove one plate of growing cells from the incubator and place it in the hood. Cells intended for transfection or other experimentation should be marked "use" during passaging, and should be placed on a different shelf from the ongoing cell lines.

3. Perform the beginning of the passaging protocol.

   1. Discard media into a waste flask.

   2. Add trypsin and allow to incubate (at room temperature) for 5 minutes.

   3. Tilt plate at a 45 degree angle and repeatedly pipette trypsin to wash cells off the plate surface.

   4. With the plate still tilted at 45 degrees, triturate the cell-trypsin solution.

      1. 1. • Note: Triturate means to slowly pipette the cells up and down to mix them into a homogeneous solution. Try not to introduce any air bubbles into the solution. Setting your P-200 (yellow) pipette to 150 µL while washing and triturating will significantly reduce air bubbles.

3. Open the hemocytometer packet and remove it.

   • Note: The following steps are for the DHC-N005 2-Chip Hemocytometer, if using a different hemocytometer, ensure that you understand the ratio of cells/volume that the average count tells you (i.e. average cells/uL or average cells/mL)

4. Add 850 uL of transfection growth media to the sterile eppendorf tube along with 150 uL of trypsinized cells (from the cell culture dish).

• This proportion can be altered, just be sure you know to what factor you are diluting your trypsinized cells.

5. Load 10 uL of the solution made in step 4 into the hemocytometer, and count all cells in the outer 4 quadrants. Use the Pre-plating Calculator tab on the Transfection Log spreadsheet. Count the cells in each box, and input that number into the green boxes at the top of the calculator. You will count four quadrants with 16 boxes each. When counting, count all cells totally within the box, as well as the cells that overlap the top and left borders of each box. Do not count cells that overlap the right or bottom borders.

   • See Hemocytometer instruction manual for insight on proper technique.

6. Change the "wells" box to set the number of wells in which you want to transfect.

7. Using the bolded and underlined values on the calculator to determine the correct amounts of each, pipette the correct volume of transfection growth media (mL media) and cell/media solution (µL cell soln) into a sterile falcon tube and vortex gently to mix.

8. Pipette 1 mL of the vortexed solution into each well and label appropriately. An appropriate label would generally include the following, as well as any other values that differ between your wells for your specific transfection:

• The name of the protein being produced in the well.

• The amount of DNA or RNA (often as a multiple of 0.5 µg, written as 1x, 2x, 5x, etc.).

• The amount of lipofectamine used (0.75 µL and 1.5 µL are common).

9. Place well tray in cell incubator and incubate for 24 hours before transfection.

Transfection

It is extremely helpful to precisely plan your transfection before performing it. Use the DNA Dose calculator on the Transfection Log to determine the amount of DNA and reagents to be added to each tube, and record the information on the Tubes tab as you calculate it. You can use the Lookup function (based on data from the Concentrations tab) and set the µg needed to calculate the amount of DNA and Lipofectamine reagent to add.

This written protocol differs from the video below. When in conflict, refer to the written protocol.

Materials Needed:

• Pre-plated cells from ~24 hours before (see Pre-plating cells for transfection)

• Lipofectamine Reagent Kit

  • • DNA kit contains two vials: Lipofectamine 3000 and Lipofectamine P300

    • RNA kit contains one vial: Lipofectamine MessengerMAX

• 0.6 mL microcentrifuge tubes

• 1000 µL, 200 µL, 10 µL, and 2.5 µL micropipettes and STERILIZED micropipette tips

• Opti-MEM

• DNA (or RNA) to be transfected

Protocol

1. You should pre-plate cells 24 hours before starting transfection. If you have not yet pre-plated cells, you will need to do so, then wait before continuing.

2. 24 hours after pre-plating, check on the pre-plated cells. They will not be very confluent, but they should appear normal.

3. Label your microcentrifuge tubes with the appropriate plasmid name and amount. It is usually helpful to arrange them in a tube rack in the same configuration as the wells, with some room leftover for the DNA tubes and lipofectamine.

4. Add 50 µL of Opti-MEM to each new microcentrifuge tube.

5. In each tube, add the predetermined amount of DNA (or RNA). In a typical transfection, add sufficient volume to contain 1 µg of DNA (or RNA).

6. In each tube, add the predetermined amount of lipofectamine P3000. In a typical transfection, this will be 2 µL.

   1. 1. 1. 1. If transfecting RNA, use MessengerMAX instead of P3000. In a typical transfection, use 5 µL.

7. Incubate 5 minutes at room temperature. During this time, centrifuge the tubes, ensuring all contents are collected at the bottom of the tube.

8. In each tube, add the predetermined amount of lipofectamine 3000. In a typical transfection, this will be 3 µL.

   1. 1. 1. 1. If transfecting RNA, do not add any lipofectamine at this step. Incubate for an additional 10 minutes as shown in step 9, but do not centrifuge a second time.

9. Incubate 10 minutes at room temperature. During this time, centrifuge the tubes again to ensure all contents are collected at the bottom of the tube.

10. From each tube, pipette 50 µL of solution into the matching well. Without letting the pipette tip touch the well contents, slowly distribute the DNA solution around the entire well.

11. Incubate the cells for 2-4 days and analyze.

You may also refer to this diagram here that outlines the transfection using lipofectamine. Our protocol is modified from this version, but it is useful in the event that our protocol does not work as expected.

Deep Freezing Cells

Materials Needed:

- Plates containing the cells desired for freezing

- Complete Growth Media

- Trypsin

- Cryostor Freezing Media

- STERILE microcentrifuge tubes

- STERILE Cryo tubes (1-2 mL capacity each)

- 1 mL and 200 uL STERILIZED micropipette tips

- Sharpie/Tool for Labeling

- Centrifuge (in back of lab)

- Mr. Frosty

- Plastic Baggies (for Day 2)

This link contains pictures of the locations of Mr. Frosty, Cryo tubes (1-2mL), and Cryostor Freezing Media:

https://drive.google.com/drive/folders/1Ilgm4j6SRzSiYEsM64yQtK70qF-7zQ3V?usp=drive_link 

Protocol

1. Aspirate media from a confluent 60 mm dish

2. Add 400 uL Trypsin to each 60 mL dish to detach cells (or 200 uL to each 35 mm dish).  Incubate for 5 minutes

3.  While waiting, set up and label sterile micro centrifuge tubes (1 tube for each dish cells), and set up and label sterile Cryo tubes (1 tube for each dish of cells)

4. Carefully wash down the cells

5. Add 1 mL RPMI 10% FBS to the trypsin and triturate

6. Transfer the liquid to the labeled sterile microcentrifuge tube

7. Centrifuge at 4,000 rpm for 5 minutes (be sure to balance centrifuge accordingly)

8. Carefully decant the supernatant without disturbing the pellet

9. Add 1 mL of fresh RPMI 10% FBS to the tube and triturate well to break up the pellet

            - Suck up the pellet with the liquid, and pipette the liquid back into the tube. Pull up on the pipette as                 you push the liquid back in (this will keep the liquid from splashing out of the tube). Repeat until                    pellet is broken up.

10. Centrifuge at 4,000 rpm for 5 minutes

11. Carefully decant the supernatant without disturbing the pellet

12. Add 700 uL of Cryostor freezing media (10 mL small glass vials in Fridge B)

13. Triturate well to break up the pellet using the same technique as in step 9.

14. Transfer the cells to the Cryo tubes

15. Place the tubes in the blue lid Mr. Frosty Container (in the cabinet above the oocyte injection station)

16. Make sure the container is filled with isopropyl alcohol

17. Take downstairs to the -80 degree C freezer and leave in the freezer for 24 hours

18. After 24 hours, remove the vials and put them in a labeled baggie (cell line, date, size of dish cells obtained from (35 or 60mm), and confluence of dish prior to freezing, etc.) and leave in the -80 degree freezer until needed for unfreezing

- Stating the size of the original dish the cells were taken from and the confluency of the dish prior to     freezing will help whoever unthaws the cells have an idea of how many dishes to divide the cells into

19. Return the Mr. Frosty to the cabinet and store upright (NOT on its side), and update the number of freezes the Mr. Frosty has undergone with the current batch of isopropyl alcohol on in the lab notebook and in the Mr. Frosty freeze count on the bottom of the protocol page.

NOTE: The Mr. Frosty needs to have its isopropyl alcohol replaced every five freezes. Do not remove the isopropyl alcohol unless the fifth freeze in a row had just occurred. Refer to the Mr. Frosty instruction manual for instruction on use and replacing isopropyl alcohol here.

Thawing Cells That Have Been Slowly Frozen in -80 °C Freezer

Materials Needed:

- Frozen Cryostor Tubes containing cells to unthaw

- Gloves to handle frozen tubes

- New, Sterile Cell culture dishes to plate frozen cells into

- Complete Growth Media

- STERILE microcentrifuge tubes

- STERILE Cryo tubes (1-2 mL capacity each)

- 1 mL and 200 uL STERILIZED micropipette tips

- Water Bath container (styrofoam box in lab)

- Styrofoam floating tube holder (in lab)

- Ethanol (at least 70%) and paper towels

- Dry Ice (preferred) or Frozen Gel Packs In Freezer B

Protocol

1. Prepare hood and sterilize hood. Be sure to have sterile pipette tips, falcon tubes, growth media, cell culture plates, and pipettes.

2. Put complete growth media in a sterile falcon tube and suspend in water bath in the back of the lab at 37 degrees C

   1. 1. 1. Use enough growth media to have at least a 1:10 ratio of cryostor media to growth media (700 uL of growth media in a tube of frozen cells, so need at least 7 mL of pre-warmed media

3. Prepare an ice bath (using dry ice/frozen gel packs in Freezer B + ice), and bring a lid.

4. Ensuring proper sterile technique (with gloves), take the cells out of the -80C freezer and place into the ice bath. QUICKLY take the cells to the pre-warmed water bath (37C) in the lab.

5. Warm cells in 37 C bath until most of the ice is gone but a small chunk remains (tube should still be cold to the touch)

   1. 1. 1. An easy way to do this is to hold the tube with your fingers and suspend in the water bath while shaking slightly every so often until a small ice chunk remains in the tube

6. Pipette frozen cell media into falcon tube with pre-warmed media

7. Mix the solution containing the cells and growth media 

8. If possible, centrifuge cells and then remove supernatant and put fresh media back in

   1. 1. 1. If not possible to centrifuge, triturate very well.

9. Evenly distribute media into the dishes

   1. 1. 1. Each 35 mm dish can hold 2.5-3 mL of media max (so 7.7 mL of media = 3-4 dishes of cells)

         2. Depending on how the density of cells you want in each dish, less media (about 2 mL each) can be dispersed among the dishes

10. Check under microscope to make sure cells are present

Where To Find Frozen Cells

The room containing the deep freezers is located on the second floor:

(Needs Reviewing)

Materials Needed:

*Note: We currently do not have any trypan Blue for staining. Many labs mix this stain with the cells to identify dead cells and exclude them from the count (blue cells = dead). For the purposes of plating an approximate number of cells in the 24 well plates for transfection the stain is not needed, but acquiring Trypan Blue in the future would be ideal for getting exact counts of cells to report publishable numbers.

• One Hemocytometer (unopened, sterile)

• 10 uL Micropipette (black with red plunger)

• Sterile 1-10 uL Micropipette Tips (red box)

• Microscope (room 515)

• Cultured cells

• Pen and Paper

• Counter or Counting App on Phone (optional, but recommended)

Counting Mammalian Cells Protocol

*IMPORTANT - For the purposes of pre-plating cells for transfection, follow the "Pre-Plating Cells for Transfection" protocol exactly as listed in order to count cells with the proper dilution. Refer to this protocol only if cells must be counted for another purpose.

1. Remove growth media from culture Cell Dishes

2. Add appropriate amount of Trypsin to dishes and incubate for 5 minutes

3. Wash cells and triturate

4. Add appropriate amount of trypsin/cell solution to sterile microcentrifuge/eppendorf tube to acquire desired dilution (ex: dilute solution by a factor of 10)

    - Diluting the trypsin/cell solution is not required, but makes counting cells MUCH easier.

5. Pipette 10 uL of triturated trypsin/cells solution and load into one side of the hemocytometer

6. If done correctly, solution should fill the selected side of the hemocytometer via capillary action

7. Place hemocytometer under microscope until 9 quadrants can be seen. Locate the four corner quadrants.

8. Select one of the four corner quadrants and begin counting the cells. Count all cells that lie within the boundaries of the overall quadrant. DO count all cells that lie on the upper and left borders of each quadrant but DO NOT count the cells that lie on the lower or right borders (image below). Record the total number of viable cells counted in each of the 4 corner quadrants and average them.

9. Multiple average cell count by the dilution factor (if diluted by a factor of 10, multiply average by 10) and 10,000

10. The result will give you the number of cells per mL of your solution that you loaded into the hemocytomer.

11. Dispose of hemocytometer once finished.

Hemocytometer Quadrant Counting Image: www.google.com/imgres?imgurl=https%3A%2F%2Fwww.hemocytometer.org%2Fwp-content%2Fuploads%2F2013%2F06%2Fhemocytometer-squares-to-count.png%3Fw%3D551&imgrefurl=https%3A%2F%2Fwww.hemocytometer.org%2Fhemocytometer-protocol%2F&tbnid=k8W3Gtgq35kibM&vet=12ahUKEwi_7r_q6Y_vAhUJbK0KHe8VA2cQMygCegUIARCoAQ..i&docid=FmnPkDGu5nEPJM&w=627&h=439&q=hemocytometer%20quadrants&client=safari&ved=2ahUKEwi_7r_q6Y_vAhUJbK0KHe8VA2cQMygCegUIARCoAQ

*Note: After determining initial density of cells in your solution using the hemocytometer, trypan blue can be used to determine % of cells that are viable. Cell viability should be above 95% for a healthy culture. This information was acquire from thermo fisher, and more information for where to order and how to use trypan blue for determining cell viability can be found at https://www.thermofisher.com/us/en/home/references/gibco-cell-culture-basics/cell-culture-protocols/trypan-blue-exclusion.html?ef_id=CjwKCAiA1eKBBhBZEiwAX3gqlyz22oBn39K7bYW_YQ5hTsg3NvFi44UOsrS9PjJfhFTKaKKz_xlOJhoCuEEQAvD_BwE:G:s&s_kwcid=AL!3652!3!305473461780!b!!g!!&cid=bid_clb_cce_r01_co_cp0000_pjt0000_bid00000_0se_gaw_dy_pur_con&s_kwcid=AL!3652!3!305473461780!b!!g!!&gclid=CjwKCAiA1eKBBhBZEiwAX3gqlyz22oBn39K7bYW_YQ5hTsg3NvFi44UOsrS9PjJfhFTKaKKz_xlOJhoCuEEQAvD_BwE 

Bulldog Bio 2-Chip Hemocytometer Instruction Manual (also found on google drive):

*Note: This protocol is for general use of the fluorescence microscope, and for viewing cells under the microscope without a microscope camera. For taking publishable-quality pictures, refer to the protocol for how to use the AmScope while following the necessary steps below to ensure the fluorescence bulb is on. The AmScope is not necessary in order to get a general idea of whether or not the cells are fluorescing, which in this case the basic viewing procedures below can be used.

Materials Needed

• Gloves

• 70% ethanol

• Cells to observe under fluorescence (either 24 well plate or cell culture dish)

• Digital Camera to take pictures of cells (if applicable)

Steps

1. Ensure the cells you will be observing have incubated in transfected eGFP (or other fluorescent protein) long enough, or ensure that the cells already have a eGFP incorporated into their genome

2. Uncover the microscope in the shared equipment room (room 515), and following sterile technique, place the dish/plates of cells to observed on the microscope stage

3. Turn on the fluorescence bulb (the green switch is on the small box to the right of the microscope), and turn the lights in the room off to make it easier to observe the fluorescing cells

   1. ***NOTE: Instructions on how to operate the fluorescent lamp are listed on the box and should be followed. Important things to remember are that the lamp should NOT be switched off until it has been on for at least 15 minutes and that the lamp should only turned on and off once per day to conserve the life of the fluorescent bulb (per Dr. Norimatsu).

4. Slide the metal knob in the center of the microscope (see video) to shine blue light on the cells if using eGFP. This microscope can shine green, blue, or both types of fluorescent light, and different types of light may be necessary depending on the pigment you are trying to fluoresce. 

5. Place the dish/plate of cells to be observed under the blue light and look into the microscope. Cells that are fluorescing should glow green (if using eGFP), while the cells that are not fluorescing should not be visibile and should just appear as black space.

6. Record any images necessary using the lab's digital camera (or a phone camera for less images of less quality)

7. Remove the cells, place back in the incubator or discard properly if no longer needed.

8. After at least 15 minutes from bulb ignition, turn off the fluorescent lamp.

Under Construction --> will need to film video describing how to use fluroescence microscope)

    - include explanations of what each knob does

    - Include what each type of light is and when it would be appropriate

    - Include example images)

Example image of LM45 cells expressing eGFP (Note: Well was near 100% confluence at time of image capture)

General

We currently have an AmScope Microscope Digital Camera that we can attach to the microscope in the shared equipment room to get a better image of our cells. In order to use this AmScope, we have a lab/cell culture laptop that has AmScope software installed on it. The AmScope must be connected to this laptop (or another computer with the AmScope camera software) in order to view the cells. The Pin for the laptop is 0504, and the password is Frog#504.

Using the AmScope for Observing Cells

1. Acquire the AmScope, lab laptop, and USB cable to connect the two (should be in the AmScope box).

2. In the shared equipment room, remove the top piece with a black chord connected to it off of the microscope. It is a cylinder-shaped piece above the eyepieces.

3. Remove the free reduction piece/adapter found within the chamber of the olympus microscope (is a black cylinder piece with lenses - says "olympus" on it.

4. Attach the AmScope reduction piece/adapter the AmScope camera body (the body is the larger square-shaped piece). 

5. Insert the connected camera body + adapter into the Olympus microscope, where the old reduction piece/adapter was removed from. Connect the AmScope body to the lab laptop via the included HDMI cable.

6. Open all shutters in the microscope, and pull the knob to switch the camera output to the AmScope. The knob to switch outputs is on the right side of the microscope, towards the base of the microscope. The main shutter you'll want to open is located on the right side of the square base the eyepiece is located on.

7. Open the AmScope application found on the lab laptop's desktop or in its applications menu.

8. If connected properly, you should see a list of options on the left of the software's menu. Under "Camera List" the MU AmScope should be listed. Click on the AmScope's name to see a live view of the camera.

To take an image/video:

1. While viewing the camera's live feed, click "capture" or "record". If recording a video, click "stop recording" when you are done filming.

2. The image or video will then appear as a new tab in the center of the screen labeled "0001, 0002, ..." etc. Rename the image/video as you see fit, and click the first tab to return to the live feed.

3. Save the images/video on the lab culture laptop or on a flashdrive. Go to file in the top left corner, click "save as" then select a location to save the image/video in. On the lab laptop, there is already a folder dedicated to AmScope images.

4. If using this saving method, each image/video will need to be saved individually. 

When observing cells for fluorescence after transfection, it is very useful to take two images (taken in exactly the same spot in the well) of everything: one under normal broad-spectrum light and one under the appropriate color of light to observe fluorescence. Between taking these two images, take care not to move the well plate or camera. You may need to refocus the microscope between images to achieve good clarity.

In the AmScope Application, the sidebar contains many settings and methods that can be used to alter how an image is viewed. It's recommended to familiarize yourself with the imaging techniques, and read about the ones you feel would be most valuable at: 

https://www.amscope.com/software/AmScope/MU-Series-Complete-Manual-Complete.pdf 

Sample Images for Reference

Images of fluorescing LM45 CD47 were taken with the AmScope on 4-9-2021. These images are available on the cell culture laptop and on the google drive for reference, and to see how some of the different filters and viewing options available can alter how the same group of cells can appear on camera.

On the google drive, the images can be found under Cell Culture Project --> AmScope Images --> Practice Images --> 04092021

On the cell culture laptop , the images can be found under File Viewer --> Documents --> AmScope Images --> Practice Images --> 04092021

After using the AmScope camera to capture images of cells, it is useful to edit them to make them easier to analyze. You can use ImageJ to do this.

1. Open ImageJ, and from the File menu, open two images: the visible spectrum image and the fluorescence image of your cells.

2. Select the fluorescence image's window. Then, under Process, select Subtract Background. Change the rolling ball radius to 100 pixels, and ensure no boxes are checked. Click OK, and the fluorescence image should become mostly, if not entirely, black.

3. Under Image > Adjust, select Brightness/Contrast. Move the brightness and contrast sliders to 80-90% of maximum. Click Apply.

4. Select the visible image's window. Under Image > Type, select 8-bit. Then, again under Image > Type, select RGB color. The image should remain in grayscale.

5. Under Image > Adjust, select Brightness/Contrast. Increase brightness and contrast only slightly to improve image clarity; the sliders should end up at 60-70% of maximum. Click OK.

6. Under Image > Overlay, select Add Image. For the image to add, select the fluorescence image. Change the opacity to 50%. Click OK.

7. Under File > Save as, select PNG. Save the file with a useful name; this would typically include the transfected plasmid, the dosage, and the date the image was taken. When finished, upload all files to the Drive and log them on the wiki.

(In progress)

Passaging

• Q: When checking my passaged cells in the days following the passaging, they always seem to be clumped in the middle of the dish.

  • A: This is presumably due to the cells not being evenly spaced out in the dish immediately following passaging. To avoid this, be sure to move the cell culture dish forward and backward AND side to side multiple times WITHOUT SWIRLING the medium. Swirling the medium causes the cells to collect in the middle of the dish, while moving in the described forward/backward and side/side motion allows the cells to evenly disperse throughout the dish prior to their adherence to the dish. If the cells are allowed to adhere to the bottom of the dish before being dispersed, they cannot be moved unless trypsin is added.

• Q: There are large streaks along the bottom of the dish where no cells seem to grow.

  • A: This are likely due to a pipette scraping along the bottom of the dish while passaging is being performed. This scraping will remove the treatment present along the bottom of the cell culture dish that allows the cells to adhere to the dish. This will not kill your cells, but it will likely prevent any cells from growing in these areas. To avoid this, refrain from touching the pipette tip to any surface of the new cell culture dish while passaging.

• Q: I am following the passaging protocol correctly and am employing proper sterile technique, but my cells still seem to be getting infected and dying.

  • A: If you are confident that your sterile technique is not the reason for your passaged cells to continue dying, ensure that the growth medium you are using for your cells is suitable. Complete growth media used for standard passaging in our lab contains RPMI 1640 + FBS as well as antibiotics. Expired antibiotics could explain the constant source of infection in your cells, but this should only be assumed when every other variable is ruled out. Be sure to record the dates, volumes, and identities of antibiotics added to growth media used for passaging.

• Q: My cells are alive, but they seem to be growing at a slower rate.

  • This could be happening for a number of reasons. As long as the cells are still behaving the same way you want them to (such as maintaining the same transfection success rate, for example), then you may not need to worry. If the rate of the cells' growth is concerning to you, you can dispose of the culture and restart your cell line by thawing out a frozen tube of cells. If this route is taken, it is recommended to make sure you successfully thaw out your chosen cell line before disposing of the old cells.

Pre-Plating Cells

• Q: Large masses appear to be floating in 24 plate wells in the day following pre-plating.

  • A: This could be due to large amounts of cells dying and floating to the surface of the wells. If this is the case, this is likely due to infection. Ensure you are practicing proper sterile technique, which can be reviewed here.

  • A: Additionally, these masses could also be large clumps of healthy cells. If this is the case, this is likely caused by an excess of cells being plated in a well on the day prior to transfection. Ensure you are staying close to the ~100,000-150,000 cells per well mark (although up to 200,000 is probably fine). These large clumps of cells are not suitable for transfection.

Transfections

• Q: I follow the transfection protocol properly, but my cells never seem to fluoresce, or very few cells do.

  • A: Remember, Lipofectamine has a relatively low success rate when it comes to actually causing the cells to express the plasmid we are trying to transfect into them. The lipofectamine producers claim about a 2% success rate, so a "successful" transfection would result in only a few cells to fluoresce. Additionally, ensure that the plasmid you are trying to transfect into the cells contains eGFP (or a fluorescent protein that can respond to the fluorescent microscope). No fluorescence will be observed if the plasmid does not have a fluorescent protein in it. All plasmid files can be found on the G drive on lab computers.

  • A: Additionally, keep in mind that transfections work best when cell lines have been passaged <50 times. For best results and success rates, use a cell line that has been passaged the fewest number of times.

  • A: Certain plasmids may be more difficult to transfect than others, such as the eGFP + Cas9 plasmid we've been practicing on. Contacting the plasmid distributor or lipofectamine manufacturer may help, or adding more DNA when attempting to transfect.

Counting Cells

• Q: There are too many cells for me to easily count (Ex: multiple hundred per quadrant). Is there an easier way to do this?

  • A: Counting multiple hundreds of cells each quadrant can get you to the same result in the end, but it's much easier to dilute your cell solution prior to counting in the hemocytometer. Diluting to a concentration that results in 10-20 cells per quadrant is significantly easier to manage. Just be sure you are aware of what magnitude you dilute your cell solution to in order to ensure you aware of the number of cells per volume in your diluted solution, as well as the stock solution of cells (from the plate). Finding a dilution ratio that is preferrable for you an the cells you are working with could take some trial and error.

• Q: I can't easily differentiate between dead and living cells in each quadrant.

  • A: Trypan blue solution can be used to alleviate this issue. Adding Trypan blue to the cells being counted allows you to differentiate between dead and living cells. Dead cells or cell fragment will stain blue, living cells will remain colorless.

(Largely needs to be edited/checked)

Passaged Cells

During passaging, old growth media should be collected in a flask and autoclaved. Once autoclaved, the old growth media can be disposed of down the drain. After old growth media is disposed of, the old cells (if healthy) can be thrown in the trash with the dish closed, after all media and trypsin is removed. It is better to remove cell culture waste often (and in small amounts if necessary) instead of waiting for large amounts to accumulate and disposing of it all at once.

It is typical in our lab currently to autoclave and dispose of old growth media after every passage.

Infection

If a plate/well gets infected, take extreme caution to isolate that plate from the rest of your cell cultures. If one plate is infected, there is a large chance your other cell lines will be infected if not taken seriously. The infected plate/well can be filled with bleach and left to sit in the fume hood (wells adjacent to the infected wells can be filled with bleach as well). After sitting for (undisclosed amount of time? Multiple minutes?) the bleach and plates can be disposed of properly.

pegFinder is a useful tool that can do large amounts of the work for us when it comes to designing pegRNA. It can be found at http://pegfinder.sidichenlab.org/.

Before designing a pegRNA, you should be able to identify the region of the genome that you would like to edit, and you should be able to access the sequence of the genome at this location. This can be done through most genome broswers, such as ensembl.

Using pegFinder:

1. Go to http://pegfinder.sidichenlab.org/

2. Find the reference sequence of the DNA you want to edit. Your reference sequence should be the DNA you are starting with, or in other words the DNA that the pegRNA will be designed to target and edit. This may be the wildtype sequence or not depending on your goal (ex: if editing mutated DNA to wildtype DNA, the mutated sequence would be your reference)

3. Enter in a desired DNA sequence, which should be the final DNA sequence you want to achieve following editing via prime editing. Make sure your reference and desired sequences align as much as possible, are no longer thatn 500 BP, and preferably contains 100 base pairs around the desired edited region of the DNA.

4. Set the desired parameters before submitting. A secondary nicking sgRNA is recommended, as it will increase the efficiency of the edit being incorporated into the genome via the prime editing mechanism. We will be using a Cas9 - NGG enzyme unless otherwise noted.

5. The results page will show multiple candidates for pegRNAs designed by pegFinder, as well as a candidate secondary sgRNA (if selected on the first page). The "full length pegRNA" consists of the generated sgF, scaffF, and entensF sequences.

After pegFinder reports pegRNA options to you, pegFinder will link to you Addgene plasmid #132777. This plasmid can be used to clone the pegRNA sequence designed, and can be ordered from addgene to perform prime edits with. (this info needs to be checked)

General

The cell incubator we will be using to store cells being used for cell culture is located in the shared equipment room on the fifth floor (room 515, right next to our lab). The cell incubator should be set to operate at 37°C, 5% CO2, and 95% O2 (conditions to mimic the human body).  If you are having issues with the cell incubator or believe it is not working properly, see Dr. Norimatsu or Blake for questions, or reference the operations manual linked below.

Cell Incubator Operations Manual

Changing CO2 Tank

If a CO2 tank needs to be replaced, call ATSU facilities (extension 2297) and have them come assist in the replacement of the tank. An old tank should be replaced when the volume of the tank is about 0, and the new tank should start at ~800 after being installed correctly. If a tank remains open consistently, it should last for about a year.

1. Ensure both your new and old CO2 tanks are within the chained region of room 515, and be sure that the metal rounded cap is covering the valve on the new CO2 tank. The chains are important to make sure a tank doesn't fall over and injure someone, therefore the tank should ALWAYS be within a chained region.

2.Turn the main valve on the old CO2 tanks off (close the valve completely).

3. Using a wrench, remove the regulator from the old CO2 tank (again, be sure the valve is completely closed before doing this).

4. Unscrew the rounded cap on the new CO2 tank and be sure the valve is closed.

5. Attach the regulator to the new CO2 tank.

6. Open the valve on the CO2 tank.

7. Adjust the pressure on the regulator to the desired range? (10-15 psi)

*** If any of these steps are unclear, PLEASE ASK someone who can help you properly replace the CO2. These tanks contain highly compressed CO2 that is flammable, can cause asphyxiation if opened for too long into the surrounding environment, or could potentially cause the tank to fly across the room if the tank falls over and the valve is forced open, therefore it should always remain chained up with the rounded cap on if not in use.

Humidity Pan

The humidity pan is the metal tray at the bottom of the cell incubator. The pan serves to maintain proper humidity levels throughout the incubator. It should remain filled with sterile distilled water at all times. The water may slowly evaporate, or the water may become contaminated over time. This could be determined by seeing things floating in the water, or by seeing growth in the water (I.e. mold, fungus, etc.). If this occurs, new distilled water must be autoclaved and used to put back in the incubator. ONLY sterile distilled water can be used, as the salts in any tap water can damage the incubator/humidity pan, and non-sterile water could lead to infection of your cells. The humidity should be filled about halfway with water at any given time, which equates to about 3L of sterile distilled water.

General

The Mr. Frosty  used for deep freezing cells should always be filled to the fill line with 100% isopropyl alcohol. The alcohol in the Mr. Frosty should be replaced every 5 freezes. After every freeze, the Freeze count should be updated to reflect the total number of freezes the Mr. Frosty has been used for since its alcohol has been replaced. After updating the freeze count, please put the date of the most recent freeze and the initials of the lab member updating the freeze count for reference if needed.

Current Freeze Count

3 freezes with current batch of isopropyl alcohol. - BS 4/23/21

The reference manual for the Mr. Frosty can be referenced at

 https://assets.thermofisher.com/TFS-Assets/LCD/manuals/Nalgene-Mr-Frosty-EN-8-0404-28-0211.pdf

 for specifics on using the Mr. Frosty or changing the alcohol.

1. FBS Aliquot

1. Slowly thaw 500 mL of FBS either overnight in a refrigerator or at room temperature until completely thawed (Note: thawing will take more than 24 hours if completely frozen)

2. Spray ethanol in hood and wipe down for sterility

3. Bring thawed FBS and 13 50 mL Falcon tubes to the hood

4. Unscrew all Falcon tube lids but DO NOT remove them

5. Transfer 40 mL of FBS from the 500 mL container to 12 individual Falcon tubes using a serological pipette attached to a manual vacuum

6. Transfer the remaining 20 mL of FBS to the 13th Falcon tube

7. Label the tubes accordingly

8. Store in -80 Freezer

2. Creation of Complete Growth Media

Note: 10 Falcon tubes of complete growth media are currently in fridge A on the middle shelf. They are contained in two separate plastic beakers labeled "RPMI 1640 + 10% FBS Complete Growth Media". Use these before making more complete growth media

1. Add 5 mL of pen/strep/amph to the 500 mL RPMI container and label

2. Add 4 mL of FBS to a sterile Falcon tube

3. Add 36 mL of RPMI + Pen/Strep/Amph to the tube containing the FBS

4. Label Accordingly

sdfasdasdfasdf

Norimatsu Platelet Count Protocol

1. Create anticoagulant solution

   1. Two separate anticoagulant solutions will be made, one containing 0.5M EDTA (pH 8) by itself, and the other containing 109mM of potassium citrate and 5mM Hepes buffer solution in a total volume of 10 mL

      1. 0.5M EDTA solution is premade and can be found in Cabinet A

      2. The second solution containing 109mM of potassium citrate can be made by pipetting 360uL of potassium citrate into 9.60 mL of water for a final volume of 10mL. To this is added 0.0119g of Hepes buffer solution

2. Obtain blood sample using Accu-Chek 

   1. Pipette (Amount) of each anticoagulant solution above to their own individual test tubes (2 tubes total)

   2. Following the directions contained in the Accu-Chek box, obtain a blood sample from the patient’s hand

      1. After puncturing the finger, use a gauze to wipe the first drop of blood

      2. Procure capillary tube and dab on the bubble of blood while simultaneously massaging it until a sufficient amount is had. DO NOT touch capillary tube to patient’s skin.  https://www.youtube.com/watch?v=ibU5PYOF2qg 

   3. Add (Amount) of blood to each of the two test tubes that contain the separate anticoagulant solutions

3. Diluting the blood sample

   1. Obtain a 20 uL sample of blood/anticoagulant 

   2. Prepare a 1.98 mL sample of dilute solution (Frog Ringer without Ca). This is a 100-fold dilution

   3. Place the blood sample in the dilution solution and mix thoroughly and allow to sit for 10 minutes

   4. Here is a video demonstrating the process: https://www.youtube.com/watch?v=_HrgH31TS7Y 

4. Examine blood under microscope

   1. Place blood/coagulant samples onto two different microscope slides

      1. The first microscope slide is a normal one, and will be streaked with the blood in the form outlined in this video: https://www.youtube.com/watch?v=4NkgEjPKzBA

      2. The second microscope slide is a hemocytometer, and blood will be added to it and the slide placed on the microscope stage for viewing. Refer to the following link for a brief protocol for using the hemocytometer: http://home.sandiego.edu/~josephprovost/Hemocytometer%20Cell%20Counting%20Protocol.pdf 

5. Count Platelets

   1. Platelet counts using a hemocytometer is demonstrated here: https://www.youtube.com/watch?v=c8WuNZh1XyA  

6. Stain Platelets (Can skip this step if viewing without stain)

   1. Dry blood smear stain

      1. Fix slide in methanol for 30-60 seconds

      2. Drain any excess methanol from the slide

      3. Flood the slide with 1 mL of Wright Stain and allow to stain for 1-2 minutes

      4. Add 1.5 mL of pH 6.8 diluted buffer (buffer type) to the stain-covered slide

         1. https://www.aatbio.com/resources/buffer-preparations-and-recipes/potassium-phosphate-ph-5-8-to-8-0 

      5. Mix the stain and buffer together by gently rocking the slide for 1 minute

      6. After rocking, allow the mixture to stand on the slide for an additional 2 minutes

      7. Rinse slide with DI water and allow to air dry

*To increase the stain strength, raise the staining time in step iii. to 3 minutes and and raise the time in step vi. to 5 minutes

1. 8. https://assets.thermofisher.com/TFS-Assets/APD/manuals/IS89013-Wright-Giemsa-Stain-Solutions.pdf 

   9. https://www.emsdiasum.com/microscopy/technical/datasheet/26060.aspx 

   10. https://theolb.readthedocs.io/en/latest/buffers/1-m-hepes-ph-70.html 

Questions:

• What kind of containers should the anticoagulant solutions be prepared in?

  • A: Microcentrifuge tubes

• Once I perform the prick of the finger and begin massaging it to get blood out, how/where do I put the blood?

  • A: Use a micropipette to remove the blood at pre-set volumes

• How much blood per anticoagulant do we want volume wise?

  • A: 100x as much blood as anticoagulant solution

• Are we going to dilute the blood? If so, in FR solution? Do the volumes I noted above look okay for dilution?

  • A: Before counting using the hemocytometer we will dilute the blood using Ca-free FR solution. A 100-fold dilution should suffice

• How much to place into hemocytometer?

  • Fill until mirror surface is just covered

  • This video also  helps answer this question: https://www.youtube.com/watch?v=pP0xERLUhyc

For the stain done on 12/21 the following protocol was followed. Some of it is taken from above, and other parts were obtained from this video: https://www.youtube.com/watch?v=9xBcm-1NMqk

1. Pipette 1 uL of ETDA per 100 uL of blood into however many micropipette tubes needed

2. Draw blood using the Accu-Chek protocol furnished above

3. Pipette the blood from finger into corresponding micropipette tube containing AC Soln.

4. Mix as well as possible and transfer blood drop onto slide

5. Perform streak as shown in the above video

6. Place the slide in methanol for 5 minutes

7. Allow to air dry

8. Add 200 uL of wright-giemsa stain to each slide and allow it to sit for 2 minutes, while shaking gently during incubation

9. Add 200 uL of DI water for 2 minutes, while shaking gently during incubation

10. Continuously rinse slides with water until the edges show a faint pinkish red

11. Dip a few times into 0.5% (1 mL acetic acid, 199 mL DI  water) acetic acid solution

12. Air dry

13. Place coverslip on slide before viewing?

(old protocol)

Complete Protocol for the Growth and Passaging of Melanoma Cells

Outline:

1. Materials Needed

2. FBS Aliquot

3. Creation of Complete Growth Media

4. Growth (old protocol)

5. Passaging (old protocol)

6. Pre-Transfection Plating (old protocol)

7. Transfection (old protocol)

1. Materials Needed and Location as of June 2019

   1. Melanoma Cells- provided by Dr. Baer’s lab

      1. Cell line 1: LM-MEL-45 (slow)

      2. Cell line 2: WM278 (fast)

   2. RPMI 1640 (Growth Media)

   3. Penicillin/Streptomycin/Amphotericin

   4. Regular FBS- -80 freezer on second floor

   5. Complete Growth Media (RPMI 1640 + 10% FBS): Fridge A, middle shelf, 10 40mL Falcon tubes 

   6. Trypsin- Fridge A, middle shelf

   7. Treated 60 mm Petri Dishes- in back cabinets with pipette tip refills, above microscope used for sorting and injecting oocytes

   8. 24 well plates

   9. Serological Pipettes and Extraction Piece to attach- in hood

   10. Access to a fume hood- Dr. Baer’s lab

       1. May be using hood in shared equipment lab in the future so we don’t keep bothering Dr. Baer

   11. Incubator (set at 37C and 5% CO2)- in Dr. Baer’s lab

   12. 0.6 mL microcentrifuge tubes*- in tube drawer

   13. All sizes of pipette tips*- various locations around our lab, make sure they’ve been autoclaved

   14. Ethanol- Fridge A

   15. Waste container

   16. Microscope (fluorescent and standard light)- Dr. Baer’s lab

   17. Desired DNA sequences- both in Freezer A

       1. eGFP

       2. eGFP + Cas9

*Must be autoclaved before use

2. FBS Aliquot

1. Slowly thaw 500 mL of FBS either overnight in a refrigerator or at room temperature until completely thawed (Note: thawing will take more than 24 hours if completely frozen)

2. Spray ethanol in hood and wipe down for sterility

3. Bring thawed FBS and 13 50 mL Falcon tubes to the hood

4. Unscrew all Falcon tube lids but DO NOT remove them

5. Transfer 40 mL of FBS from the 500 mL container to 12 individual Falcon tubes using a serological pipette attached to a manual vacuum

6. Transfer the remaining 20 mL of FBS to the 13th Falcon tube

7. Label the tubes accordingly

8. Store in -80 Freezer

3. Creation of Complete Growth Media

Note: 10 Falcon tubes of complete growth media are currently in fridge A on the middle shelf. They are contained in two separate plastic beakers labeled "RPMI 1640 + 10% FBS Complete Growth Media". Use these before making more complete growth media

1. Add 5 mL of pen/strep/amph to the 500 mL RPMI container and label

2. Add 4 mL of FBS to a sterile Falcon tube

3. Add 36 mL of RPMI + Pen/Strep/Amph to the tube containing the FBS

4. Label Accordingly

4. Growth

   1. 4 days to 1 week at a time

5. Passaging

1. Wash hands and put on gloves, spray gloves with ethanol.

2. Turn on the blower and light for the hood and open it

3. Clean the hood with ethanol, wiping from back to front

4. Procure complete growth media (RPMI 1640 + 10% FBS) and trypsin from fridge and place in hood

5. Place previously autoclaved micropipette tips of all three sizes in hood along with the pipettes

6. Unscrew the caps to the media and trypsin but leave them on 

7. Obtain cells from the incubator and place in the hood 

8. Check viability of each dish of cells by looking under microscope

   1. Cells should be 100% confluent

   2. Refer to Photos of 100% Confluent Melanoma Cells Before Passaging

9. Place new cell dishes in hood and label with name and date

10. Remove the lid of the dish that currently contains cells and place lid facing down, tilt dish at a 45° angle to have the media collect in one spot.

11. Pipette out the media from each dish using the suction pipette/aspirator, made sure to not let the pipet tip touch anything but the plate (not your gloves) and use a new pipet tip for each plate.

12. Add the appropriate amount of trypsin to each dish 

    1. 400 uL to 60 mm dish 

    2. 30 uL to 35 mm dish

13. Allow the cells to incubate in the trypsin in the hood for 5 minutes

14. During this incubation period, add the appropriate amount of complete growth media to the new dishes 

    1. 4 mL to 60 mm dish

    2. 2 mL to 35 mm dish

15. After incubation period, angle the parent dish at 45o and suction trypsin using a micropipette and wash the cells down the side to diminish any remaining adherence

16. Triturate to mix the cells

Note: triturate means to slowly pipette the cells up and down to mix them into a homogeneous solution, try not  to introduce any air bubbles into the solution

1. Add two drops (or more if needed) from the parent plate to the new plate

2. Spread the cells all over the new plate by pipetting up and down

3. Gently move the plate from side to side, up and down, to evenly spread the cells

   1. Do not move the plate in a circular motion or the cells will collect in the middle of the dish and will not be evenly spread out.

4. Check the new plate under a microscope to ensure the cell amount and distribution is sufficient

5. Discard the old cells in the trash

6. Pre-transfection plating

1. 1 day before transfection, remove the plates that contain the growing cells from the incubator and place them in the hood

2. Follow the passaging protocol as normally done:

   1. Discard media

   2. Add trypsin and allow to incubate (at RT) for 5 minutes

   3. Tilt plate at a 45 degree angle and pipette trypsin to wash cells off the plate surface over and over

   4. With the plate still tilted at 45 degrees, triturate the cell-trypsin solution and then allow to sit

Note: triturate means to slowly pipette the cells up and down to mix them into a homogeneous solution, try not  to introduce any air bubbles into the solution

3. Procure hemocytometer, microscope, and one sterile falcon tube as well as a sterile 1 mL (or greater) microcentrifuge/eppendorf tube and click counter to count the cells

   1. NOTE: The following steps are for the DHC-N005 2-Chip Hemocytometer, if using a different hemocytometer, ensure that you understand the ratio of cells/volume that the average count tells you (i.e. average cells/uL or average cells/mL)

4. Add 850 uL of growth media to the sterile microcentrifuge/eppendorf tube along with 150 uL of trypsinized cells (from the cell culture dish).

5. Load 10 uL of the solution made in step 4 into the hemocytometer, and count all cells in the outer 4 quadrants using proper technique (only count cells that line on the borders of the upper and left border. DO NOT count cells that lie on the borders of the lower and right border).

   1. See Hemocytometer instruction manual for insight on proper technique

6. Calculate the average number of cells from these four values. This value is the number of cells per volume (number of cells/mL of solution used for counting) of the cell/media solution made in step 4.

7. Divide the average by 1000 to find the number of cells per uL.

8. Calculate the amount of uL needed of the cell/media solution prepared in step 4 to plate the desired number of cells per well (100,000-150,000 cells/well). For example, if you want 100,000 cells per well and you’re plating 4 wells, take 400,000 divided by the number of cells/uL. This value will give the volume in uL of cell/media solution needed to have 400,000 cells.

9. Calculate the amount of growth media needed to create a solution of growth media with cells equating to the desired number of cells per well (such as 100,000) with each well containing 1 mL of solution. First, account for 1 mL of growth media for each well desired for plating. (ex: 4 wells → 4 mL of growth media).Take the total volume of growth media and subtract it by the volume of cell/media solution calculated in step 15. For example, if plating 4 wells and you calculated 200 uL of cell/media solution needed for 400,000 cells (when wanting 100,000 cells/well), you would do 4 -.2 to get 3.8 mL of growth media needed.

10. Pipette the correct volume of growth media and cell/media solution into a sterile falcon tube and vortex gently to mix.

11. Plate 1 mL of the vortexed solution per well and label appropriately.

12. Incubate for 24 hours before transfection.

13. Allow to incubate overnight before transfection

7. Transfection

*DON’T FORGET to autoclave 0.6 mL eppendorf tubes before the start of the transfection process, as some will be needed

1. On day 1, plate melanoma cells at 100,000 cells per well (count this using the hemocytometer) and label two wells as 0.75 and 1.5. Refer to protocol above for this step.

2. On day 2, check on the melanoma cells and ensure confluency is about 70% (subjective viewing under a microscope)

3. Label 3 0.6 mL eppendorf tubes as 0.75, 1.5, and DNA

4. Add 25 uL of serum free media to the eppendorfs labeled “0.75” and “1.5”

5. Add 50 uL of serum free media to the eppendorf labeled “DNA”

6. Add 0.75 uL of the lipofectamine 3000 reagent to the eppendorf tube labeled “0.75”

7. Flick the eppendorf tube about 5 times to ensure it is well mixed

8. Add 1.5 uL of the lipofectamine 3000 reagent to the eppendorf labeled “1.5”

9. Flick the eppendorf tube about 5 times to ensure it is well mixed

10. Add 1 ug of DNA to the eppendorf labeled “DNA” 

11. Add 2 uL of lipofectamine P3000 reagent to the eppendorf labeled “DNA”

12. Flick the eppendorf tube about 5 times to ensure it is well mixed

13. Add 25 uL of the serum-free media, DNA, and P3000 mixture (from the DNA eppendorf) to the eppendorf labeled “0.75”

14. Add 25 uL of the serum-free media, DNA, and P3000 mixture (from the DNA eppendorf) to the eppendorf labeled “1.5”

15. Let the mixtures incubate for 5 minutes

16. Add 50 uL of the DNA mixture from the eppendorf labeled “0.75” to the 0.75 well on the 24 well plate. When adding the DNA, pipet slowly and in a circle to mix the DNA into the media.

17. Add 50 uL of the DNA mixture from the eppendorf labeled “1.5” to the 1.5 well on the 24 well plate. When adding the DNA, pipet slowly and in a circle to mix the DNA into the media.

18. Incubate the cells for 2-4 days and analyze

You may also refer to the diagram HERE that outlines the transfection using lipofectamine.