Month / Mar 2023

by Gabriele Feigelman

Mouse Tail Vein Injection Protocol

List of Equipment:

  • Syringes (for injection and preparation)
  • 18-19 G needle (for preparation)
  • 27 G needles (for injection)
  • Vortex
  • Saline (100μL/250,000 cells)
  • Infrared (IR) lamp (for warming up mice)
  • Empty cage (for transferring mice after injection)
  • Mouse restrainer
  • Marker
  • Follow-up pages (for recording injection volume, weight, and other details)
  • Scales
  • Pad gauze
  • Alcohol (70%) for hand hygiene

Procedure:

  1. Weigh the mouse and fill out the necessary paperwork.
  2. Warm up the mice for 10 minutes by exposing them to an infrared (IR) lamp from 10 cm. A Pro Tip: This step is critical because proper preparation leads to success! Do not forget to remove the green lid and the housing from the cage)
  3. Vortex the tubes with the cell suspension and use an 18-19G needle to collect the fluid. Change to a 27G injection needle. Discard any excess back into the tube so that just 100μL would stay in the syringe. NB. Check for bubbles and remove any excess fluid, as this can help verify the plunger’s ease of use.
  4. Align the tip of the 27G needle with the scale on the syringe facing upwards.
  5. Place the restrainer on the table border, ensuring that the border of the restrainer is aligned with the table border.
  6. Transfer the mouse tail-first to the restrainer. Keep the mouse by tail basis firmly and locate the side veins.
  7. Move your fingers down the tail slightly, but maintain tension to uncover the first third of the tail near the basis.
  8. Align the needle with the syringe so the tip hole is parallel to the tail, facing upwards (180°).
  9. Insert the needle into the vein, ensuring that it goes in at least 75% of the way to make it stable.
  10. Gently pull the plunger backward to observe a flashback of blood. If no flashback is observed, try moving the needle back and forth until a flashback is seen.
    Pro Tip: Only inject the content of the syringe after confirming the flashback.
  11. After confirming the flashback, dispense the content of the syringe into the vein.
  12. Apply pressure with a pad gauze to prevent bleeding while marking the mouse with a marker. Then, place the mouse in an empty follow-up cage.

A Pro Tips:

  • Based on my experience, if you’re working with mice, it is essential to have a reliable restrainer to ensure safety, comfort, and successful technique. I highly recommend the BrainTree Tailveiner® Restrainer for Mice (TV-150 STD). In my opinion, it’s the best option out there, and it’s my personal favorite. Make sure to use a good restrainer, as it’s the most crucial piece of equipment you’ll need.
  • Before starting, ensure that the mouse is properly warmed up. Check heating lamps before because they differ by their heating strength.
  • Again, inject after confirming the flashback with blood in the syringe/needle.
by Gabriele Feigelman

Protocol for Decellularization of Deposited Lung ECM from Primary Lung Cells

  1. Preparation of Lung ECM:
  • Obtain fresh mouse lung tissue and rinse with PBS to remove blood or debris by perfusing 3 ml PBS through the trachea or main bronchi.
  • Place the lung tissue on a cell strainer 70 microns on a 50 ml falcon tube.
  • Gently mash them with a plunger to extract the cells and tissue fragments.
  • Rinse the cell strainer with sterile PBS to collect any remaining tissue pieces.
    A Pro tip: Lift the cell strainer a bit from time to time to improve PBS drainage.
  • Centrifuge the collected cells at 2000 RPM for 7 minutes to remove red blood cells.
  • Implement red blood cell lysis if needed (Sigma Product No. R 7757):
  • Add 1 ml of buffer to the cell pellet.
    • Gently mix for 1 minute.
    • Dilute the buffer with 15 ml of PBS.
    • Centrifuge at 2000 RPM for 7 minutes and decant the supernatant.
  • Resuspend the cell pellet in full media (DMEM, 10% FCS, 0.5 % Pen-Strep, 1% glutamine, 1% NEAA, 1% pyruvate,1% fungizone) at a density of 200,000 cells per 75 microliters with 37.5 microliters of supernatant from HL-60 cells that are enriched with growth factors.
  • Plate the lung cells onto a 96-well plate and incubate for 72 hours to give time for cells to attach and secrete the ECM.
  • Preparation of Lung ECM:
  • After 72 hours of incubation, remove the media from each well.
  • Wash each well with 100 microliters of sterile PBS to remove any cellular debris.
  • Place 100 microliters of sterile distilled water (DDW) into each well to decellularize the wells for 30 minutes.
  • Remove the DDW and add 37.5 microliters of fresh ______ full media to each well. (depending on the cell line)
  • Seeding of Cells:
  • Prepare a cell suspension of 20,000 cells in 37.5 microliters of ____ full media. (depending on the cell line)
  • Plate the cell suspension in quadruplicates on a 96-well plate containing either lung-decellularized ECM or plastic.
  • Switch  ________ full media to serum-free media with or without treatments after overnight incubation. (depending on the cell line)
  • Incubate the plate at 37°C and 5% CO2 for ____ hours. Recommendation (48-72 hours)
  • Analysis:
  • After the 96-hour incubation period, assess the cell viability and proliferation using standard assays such as CCK-8, and collect supernatants.
  • Compare the results of the cells cultured on lung decellularized ECM with those cultured on plastic to determine any differences in cell behavior.

For 96-well plates, you can download a protocol template:

protocol-for-decellularization-of-deposited-lung-ecm-from-primary-lung-cellsDownload
by Gabriele Feigelman

Protocol for Orthotopic Mouse Tumor Injection (Breast Cancer):

Materials:

  • Cultured tumor cells (200,000 cells in 40 µL saline per mouse)
  • 1.5 mL Eppendorf tubes
  • Sterile 27-32 G needles (A Pro tip: Adjust needle size by checking if counted cells stay alive in vitro after drawing them and dispensing in plate vs. the exact number routinely seeded – if not, lower the G of your needle)
  • 1 mL syringe
  • Isoflurane
  • Q-tips (used for iodine solution and eye cream applications; to give an extra stretch to the skin)
  • Iodine solution
  • Balb/c mice
  • An eye cream that prevents dryness
  • Gauze Pads

Preparation of Cells:

  1. Mix the tumor cell suspension well but gently.
  2. Draw the cells without the needle attached.
  3. Attach the needle (to remove bubbles) and adjust the tip to align with the scale.
    A Pro tip: If you want to eliminate bubbles, draw more L than you need. You’ll need to draw some air into the syringe to increase the distance between the fluid level and the tip. To remove bubbles, flick the syringe. Empty all air and dead space until you see the first drop at the tip of the needle. Make sure you leave only the volume necessary for injection.

Preparation of Mice:

  1. Weigh and number the mice.
  2. Anesthetize the mice with isoflurane.
  3. Cover the eyes of the mice with protective ointment to prevent dry eyes.
  4. Lay the mouse on the gauze pad before shaving.
    A Pro tip: It is easier to collect hair with gauze pad.
  5. Shave the mice.
  6. Remove all the hair with the gauze pad.
  7. With a Q-tip dipped in an iodine solution, scrub the injection area once.
    A Pro tip: Iodine solution stains all the skin except the nipple area, which makes it easier to identify injection sites.

Injection Technique:

  1. Identify the fourth abdominal mammary gland.
  2. Lift the skin (3-4 mm) near the fourth mammary gland nipple area while placing the forceps (1-2 mm above the nipple) before injecting.
    A Pro tip: Some mice may have stretchy skin that obstructs vision. To improve access, ask your colleague to gently stretch the skin above and below the injection site using Q-tips.
  3. Insert the needle (2-3 mm) from the side at an angle of 180° under the nipple area. The tip should be inserted with an additional 0.5-1 mm.
    A Pro tip: Once the needle has passed the skin barrier and been positioned at the correct depth, lift the tip of the needle with the nipple. The cells are injected beneath the nipple.
  4. Inject 40 µL (not more than 50 µL) of the tumor cell suspension.
  5. Remove the needle slowly to avoid leakage.
  6. Injecting properly will form a visible, palpable round bump that disappears without leaking any cells.
  7. Observe the animal after waking up from anesthesia in its follow-up cage.
  8. Return the animal to its original cage.

    Tumor growth and development should be monitored twice a week in the mice. Depending on the cell type and number injected, tumors are palpable approximately seven days after injection and can be measured with calipers. It should take 2-3 weeks for tumors to form a tumor that can be removed easily.

    It is highly recommended that you watch this video, even though the injection technique is different, in order to see how the fourth mammary gland nipple area is identified and what the bump looks like.
by Gabriele Feigelman

Bradford Assay

Materials:

  • BSA (bovine serum albumin) standard solution stored with a concentration of 1.4 mg/ml
  • Bradford reagent (dye-binding reagent)
  • Test samples containing protein
  • 96 well Microplate
  • Spectrophotometer

Procedure:

  • Prepare the BSA standard solutions of different concentrations (e.g., 0, 0.05, 0.1, 0.2, 0.3, 0.4, 0.5, and 0.6 µg/µL) by diluting the BSA stock solution with distilled water as follows.

    BSA standard preparation for working concentration 1mg/ml
  • 45 µL (BSA 1.4 mg/ml in DDW) +18 µL DDW
Final BSA Concentration µg/µLVolume BSA from 1 mg/ml (µL)Volume DDW (µL)
0025
0.051.2523.75
0.12.522.5
0.2520
0.37.517.5
0.41015
0.512.512.5
0.61510
  • Add 10 µL of each BSA standard solution to the 96-well microplate in duplicates.
  • Lysates diluted 1:10 in DDW, e.g., 3 µL Lysate + 27 µL DDW.
  • Add 10 µL of each diluted lysate in duplicates.
  • Add 150 µL of Bradford reagent using a multi-channel pipette.
  • Measure Immediately OD at 595 nm using a spectrophotometer.

PRO TIP: Practice proper pipetting technique as it is clearly presented here.

by Gabriele Feigelman

How to Prepare Lung Lysates from Mice Lung Tissue

  1. Label Eppendorf tubes.
  2. Prepare the Lysis Buffer by mixing 270 µL with 30 µL of Protease inhibitors (Sigma Cat. No S8820, 10x) for one lung tissue measuring approximately 2×2 mm. The ratio should be 1:10. Calculate the required volumes based on the number of specimens:
  • N of specimens x 300 µL = Final Volume
  • N of specimens x 30 µL = Required volume of protease inhibitors
  • Final Volume PBS = Final volume – Required volume of protease inhibitors
  1. Sonicate specimens on ice using short pulses not exceeding 5 seconds.
  2. Incubate specimens on ice for 30 minutes to cool down.
  3. Centrifuge at 12,000 RPM for 15 minutes.
  4. Collect fresh supernatant in a separate Eppendorf tube.
  5. Determine the protein concentration using the Bradford Assay (see Protocol for the Bradford Assay.

    PRO TIP: Maintain optimal cooling conditions during lung tissue preparation, sonication, centrifugation, and supernatant collection