BAL FOR ADENOSINE LEVELS, CELL COUNTS AND MEDIATORS
The key to this protocol is working quickly to collect and process the samples. Use ice cold reagents and process each mouse individually in order to more rapidly freeze down the BAL for adenosine levels.
Nucleoside Preservation Cocktail (NPC) to prevent adenosine degradation: The stock solution (1000X) is 10 mM; working solutions (1X) are 10 µM. Store stock at -20°C; working solutions at 4°C. Working solution is 10 µM dipyridamole, 10 µM ab-methylene ADP and 10 µM 5’-deoxycoformycin (DCF). This will be provided by the Blackburn Lab.
Make working dilution by adding 20 ul of stock to 20 mls PBS. Keep cold. This is what you will lavage the mice with.
1. Lightly anesthetize animal with Avertin (2.5% working solution in sterile PBS), or you can use whatever anesthetic you are use to using.
2. Using blunt dissection, expose trachea, cut into trachea, and secure a blunted 21 gauge needle into the trachea using thread.
3. Slowly infuse 500 ml (depending on age of animal) NPC-PBS into the lungs using a syringe. Next, slowly draw out the NPC-PBS taking care not to pull out the needle. This can be done by holding the needle in place while extracting the BAL fluid. Remove the syringe from the needle and expel the contents into a 15 cc conical tube on ice. Repeat this 3 times, collecting 1 to 2 ml of BAL fluid. Keep this on ice.
4. Mix BALF gently and remove 20 ml of BALF and place in a microfuge tube on ice. Use this aliquot to count total BAL cells using a hemocytometer.
5. Centrifuge the BAL fluid at 1,200 rpm at 4°C for 5 min.
6. Determine and record the total volume of the BAL fluid.
7. Gently remove the supernatant without disturbing the cellular pellet. Aliquot into three separate tubes and then flash freeze in liquid nitrogen. You can also flash freeze the remaining pellet.
Important: Process each mouse individually all the way through before going to the next mouse to insure the BAL is frozen as rapidly as possible. Work quickly but carefully.
PROTOCOL FOR INTRATRACHEAL (I.T.) BLEOMYCIN CHALLENGE
Normal saline (or PBS), sterile filtered
Teva Pharm or from other clinical grade supplier, Lyophilized powder in sealed glass container (15 or 30U).
Use disposable 1ml syringe with needle to add enough sterile saline (0.5 or 1ml) to obtain a 0.03U/uL dilution. Distribute 25 or 50 ul aliquots to plastic tubes that can be stored at -80°C for a few months.
Dose: 2.5 U/kg/50 ul, 0.03 U/20 g mice. Mice should be 8-10 weeks old.
1 mouse: 2.3 ul of the 0.03U/ul stock + 47.7 ul st. saline
Prepare fresh solution every time from freshly thawed aliquot.
Control mice receive 50 ul sterile saline.
Avertin: 100% stock, 2.5% dilution in saline, sterile filtered (protected from light with foil or black tubes). Avertin stock should not be older than 1 month. Prepare the diluted Avertin the day of the experiment or a day before the experiment at the most.
Surgiloc S/C tissue glue (Fischer Scientific)
Betadine or Chlorhexidine
Marcaine 0.25%, Abbott Laboratories
Dissection scissors, straight
2 pairs of forceps: one curved and one straight
Clippers (sideburn or beard trimmers work well)
Hamilton 710 LT syringe 100 ul, 80601 and needle N726S NDL (26s/2”/2)
Disposable 1mL tuberculin syringe with needle
Incandescent lamp or heating pad and a 20-30 degree inclined plane
MAKE SURE EXPIRATION DATES OF ALL CONSUMABLES ARE CURRENT
The surgeon should wear cap, mask, clean lab coat or gown, and sterile surgeon’s gloves
1- Before starting the procedure
Bring previously autoclaved surgical tools and prepare sterile solutions of Avertin, Bleomycin, and PBS on the day of the experiment.
2- Prepare surgical area
Designate sterile surgical and recovery areas. Prepare the areas by spraying the surgical and recovery areas with at least TWO hard surface disinfectants such as Isopropyl alcohol (70-99%), Chlorine Dioxide (Clidox), Phenolics (Lysol) or Chlorhexidine (Nolvasan). Wait a few minutes (2-3) for the disinfectant to have an effect and wipe off the surgical area, clearing any debris that may be present.
Make sure that you also disinfect any surgical equipment, such as support tables that will be in close proximity with the animal.
The recovery area must include an inclined plane (20-30 degrees) and a heating lamp or pad.
3- Prepare mouse for operation
Anaesthetize mouse with 2.5% Avertin (250-300ul for 20g mice) injected i.p. using the tuberculin syringe. The animal is properly anaesthetized when pinching the hind foot elicits no response.
Place the mouse on its back and using the clippers, carefully trim away fur from the surgical area (throat).
Disinfect by applying first isopropyl alcohol on the surgical site, repeat the process using betadine, and finally ethanol to clean the betadine from the surgical site (betadine is an irritant).
Marcaine (bupivacaine, 0.25%) is injected under the skin (SQ). Inject small volumes of Marcaine around the site of incision. Do not administer more than 1ml/kg (20ul for a 20g mouse). Wait a few minutes (3-5) before starting the surgery. Marcaine lasts for 10-12hrs in the mouse.
4- Surgical Procedure
Before starting the surgery ensure that the mouse does not react when you pinch its hind foot and that Marcaine has been adequately administered to the animal.
Make a small (~1cm) incision with fine scissors and expose the trachea by blunt dissection using the forceps, taking care not to damage the surrounding tissue. Place the curved forceps under the trachea to hold it.
Gently push the needle of the Hamilton syringe into the trachea just below the larynx through the muscle and connective tissue. You only need to insert a few mm of the needle.
Slowly insert 50 ul of the solution to be administered, while the trachea and the syringe are kept steadily in place. A few seconds later pull the needle straight back, the trachea will reseal automatically.
Hold the mouse up for a few minutes by grabbing the fur at the back to help proper breathing.
5- Resealing the incision
Place the mouse on its back again and align the sides of the wound, taking care not to get fur inside.
Apply a few drops of Surgiloc to glue the skin together.
Place the mouse on the elevated pad under a heating lamp until it recovers from anesthesia. To ensure proper heating and drug distribution, turn the mouse from one side to the other every 5 minutes. Return mice to their cages when they are able to move around. The cages should be clean with fresh bedding and have enough drinking water and moist food. Include a post-op label on the cage and monitor them daily for 3 days after surgery, and then less frequently until the end of the 14 day period, when samples are collected.
7- Procedures on multiple mice
After performing the procedure with the first mouse, place tools in the bead sterilizer for at least 5 minutes, followed by 15 minutes to cool off with 70% ethanol. During this time you can change your gloves to new sterilized gloves. Also make sure to sterilize the Hamilton syringe by letting it sit in 70% ethanol for at least 15 minutes. Do not use 100% ethanol as it will evaporate too fast, precluding effective sterilization. Repeat this procedure for as many mice are needed in the study.
PROTOCOL FOR SHIPPING SAMPLES TO THE BLACKBURN LAB
1- Please label the tubes with box letters and in clear, easy to read writing using a permanent marker.
2- Label the top and side of the tube to ensure that the sample will be identifiable.
1- Place sample tubes in a CARDBOARD freezer sample box WITH NO HOLES containing tube separators (do not use plastic boxes) and ensure that the box is completely and securely closed.
2- Inside the box, include a list of the samples that are packaged in the box
For example: LUNG samples no. L1-L10; BAL fluid no. B1-B10; Serum samples S1-S10
3- If sending more than 1 box, label each box clearly with 1, 2, 3 etc.
4- Secure the box further by wrapping rubber band across the box; use at least 2 rubber bands. Ensure that the box has NO HOLES to prevent samples from falling out of the box and subsequent loss of samples.
5- Under no circumstances send samples in plastic bags or in uncovered containers; fastening tubes to an uncovered rack will lead to all of the samples coming loose during transport and loss of sample.
1- Find a suitable polystyrene box for shipment of the samples and a suitably sized cardboard box that fits the polystyrene box.
2- Create a layer of dry ice at the bottom of the polystyrene box.
3- Place the freezer boxes containing your samples roughly in the middle of the polystyrene box and fill with dry ice, adding it to the top and sides.
4- Make sure you can securely close the lid of the polystyrene box.
5- Insert the polystyrene box inside a regular cardboard box for shipment.
6- Remove, blackout, or cover ALL previous addresses on the cardboard box to prevent accidental delivery of the box to an unwanted address.
7- Place shipment IDEALLY on Monday or Tuesday to ensure the samples do not sit over the weekend in a hot warehouse.
8- Please notify Kelly Volcik (primary contact), Harry Karmouty (secondary contact), and/or Ning Chen (secondary contact) before shipping the samples to confirm that we will be able to receive the shipment on the expected delivery date. [contact information is provided at the bottom of this page]
9- Shipping address:
UTHSC – Medical School
Attn: Blackburn Lab, MSB 6.200
Houston, Tx 77030
NOTIFICATION OF SHIPMENT
2- In your email please include:
Number of sample boxes being sent
Detailed sample information
Example: BOX1 Lung BAL samples from hypoxia and LPS models, 40 samples total; BOX2 Kidney samples from KO mice, 33 samples total
3- Please include the TRACKING number of the shipment.
RECEPTION OF SAMPLES
1- As soon as we have received, stored, and checked for the delivery items, Kelly, Harry or Ning will email the person who notified us of the delivery.
2- We will confirm that your shipment has arrived and notify you of any discrepancy regarding the number of sample boxes or actual samples received.
- The main lab number is 713.500.6047
- Kelly’s contact number is 713.500.7886 (office)
- Harry’s contact number is 832.655.7678