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    • CommentAuthormaangel
    • CommentTimeMay 15th 2012
     

    GENERAL MAINTANENCE
     

    Needle-Making

    NOTE: Use the glass tubes with filament (World Precision Instruments, Inc; 1.0 mm; 4 in). Make needles BEFORE the morning of microinjection. 

    1) Turn the needle-pulling machine on.

    2) Place 1 glass tube through the black apparatus at the top of the needle-pulling machine and tighten.

    3) Drag the moving part of the black apparatus up until it is close to its stopping point and tighten. The middle of the glass should be between the top and bottom parts of the apparatus. 

    NOTE: When pulling needles, I set all the parameters of the needle-pulling machine at 990. However, these can be changed at your own convenience.

    4) Hit the “Pull” button on the bottom right hand corner. The coil should burn bright red (BE CAREFUL!) and the glass should pull, creating two needles.

    5) After the coil has cooled, carefully remove the newly created needles. Avoid touching the tips of the needles.

    6) Repeat for as many needles as you want/need. (I would recommend making 10-15). 

    NOTE: The following instructions are what worked best FOR ME. This may change for you. Find the method that works best for you and use that; no one way exists to do this.

    7) Examine the tips of the needles under the microscope. There should be a microscopic hole at the end, barely large enough to see. If you can instantly see the hole, it is most likely too big. If you determine a hole must be made, follow the instructions below. If not, store the needle for future use.

    8) Place the needle in the magnetic needle holder connected to the Picospritzer and focus on the very tip of the needle. 

    9) Using fine forceps, VERY gently scrape the edge of the needle in an up-down motion to create a tiny hole. The hole you make should be smaller than you think you are going to need. 

    NOTE: You can also use the yellow “sandpaper” to create needles, although I found that method to be too variable.
    10) Make 10-15 needles so you are prepared for future microinjections. Store them by securely placing them in clay in a large petri dish.
     
    Microinjection Plate (taken from Lom Lab Protocols)
    1) Boil (by using microwave) 3% agar into solution (between 30-50 ml). DO NOT PLACE THE MOLD INTO BOILING AGAROSE DIRECTLY OUT OF THE MICROWAVE. Extreme heat causes warping of mold.

    2) Cool agarose to 45ºC on bench top. Use a digital thermometer to monitor the temperature.

    3) Attach a tabbed piece of tape to the backside of the dried mold. This acts as
    a handle to control the mold plate as it enters the agarose.

    4) Pour the cooled agarose (45ºC) into a Petri dish.

    5) Allow one end of the mold to contact liquid, then lay the mold on top of the
    liquid in one smooth motion. This “floating action” eliminates air bubbles.

    6)  Once the mold is in the liquid agarose, allow it to solidify at room
    temperature. This should take ~20 minutes to completely solidify.

    7) Place the agarose petri dish that has firmed to an opaque color into a 4ºC
    refrigerator for approximately 30 minutes.

    8) Slip a spatula gently under the perimeter of the mold to assist separation
    from the agarose.

    9) Slowly pop the mold out. If done properly, the mold should release cleanly.

    10) The gel injection plate you just made is reusable. You will most likely have to remake the injection plate at least once in a semester, although that is dependent on how carefully you handle the embryos embedded in it. To store, flood the impression with same liquid used to make the gel, cover the dish, and refrigerate.
     

    Day before microinjection
     

    Pair Fish

    Between 4:30-8 PM (before lights are turned off), pair fish in system water in small bins on the cart, using the divider to separate the two females and one male. Leave these fish overnight.

    Fill Air Tank

    Make sure the air tank is filled. If it is not, follow the instructions below.

    1) Disconnect the air tank from the black cord connecting it to the Picospritzer using the wrench.

    2) Roll tank to the hood using a tall chair.

    3) Fill the air tank by placing the air line (the blue cord) over the metal part of the tank with the red knob. 

    4) Fill the tank with air until you can no longer hear air rushing into the tank.

    5) Reconnect the air tank to the black cord and thus, the Picospritzer.

    Day of microinjection
     Mate Fish

    1) As soon as the light comes on (around 8 AM, although this changes), move the bins to the well-lit, temperature-controlled lab room.

    2) Remove the dividers from the bins.

    3) About 45 minutes later, check for eggs. If eggs are present, return the fish to the vivarium, collect the eggs in a mesh net, and place them in a large plastic cup found in the lab room. If eggs are not present, return every 15 minutes until eggs are present. Do not collect eggs past 10 AM because they are often not fertilized.

    NOTE: The eggs must be microinjected at the 1-2 cell stage, so make sure to collect the eggs soon after they have dropped. You may have to check the bins every 10-15 minutes to assure properly aged fish.

    Microinjection
     

    1) Load the needle with ~4ul of the appropriate solution using the extra long pipette tips.

    NOTE: The following instructions are for the Chd-MO, a control morpholino with a known phenotype (see Nasevicius & Ekker 2000), that we use as an estimate for Slitrk-MO toxicity. It can be ordered from GeneTools (https://store.gene-tools.com/Standard/VIEWCATEGORY/UHJlcGFyZWQgQ29udHJvbCBPbGlnb3M%3D/) and stored at room temperature in the bottle in which it came. 

    2) I diluted the Chd-MO to concentrations based off those used in Nasevicius and Ekker (2000), which first characterized the Chd-MO phenotype. The calculations below rendered the two working concentrations I used in my experiments—0.08 ng/nl (1:100 dilution) and 0.8 ng/nl (1:10 dilution). Because I was not able to fully determine the proper Chd-MO concentration, begin with these two concentrations, understanding they might need to be altered.

     

    STOCK1: 1 mM in H2O (8 ug/ul)
     
    WORKING #1: 0.08 ng/nl (1:100 dilution)
                1:100 dilution in 4 ul TV = 0.04 ul of the stock (unpipetteable)
               
    SO, take 1 ul of stock + 9 ul of H2O (1:10 dilution, which is your stock2)
    THEN, use 0.4 ul of 1:10 stock2 + 3.6 ul of E3/phenol red (TV=4 ul)
     
    WORKING #2: 0.8 ng/nl (1:10 dilution)
                0.4 ul of STOCK1 + 3.6 ul of E3/phenol red (TV=4 ul)
     

     

    3) Insert the needle into the needle holder and tighten the screw to hold it in place.
    4) Adjust the microinjection device to the position that is most comfortable for YOU to inject.
    5) Make sure that all devices (Picospritzer, needle, air tank, etc.) are connected properly.
    6) Pipette the eggs onto the microinjection plate.
    7) Align the eggs in the rows on the microinjection plate using the fine forceps. Be careful not to rip them.
    8) Carefully remove some of the liquid using a pipette from the microinjection plate, but leave enough so that the eggs are barely covered. If there is not enough E3 in the plate, the eggs will dry out and die.
    9) Adjust the Picospritzer to the appropriate “Duration” (generally 10-20) and test the amount of liquid ejected by pressing the foot pedal. Only a small amount of liquid should be released from the needle.
    NOTE: If too much liquid is released, decrease the duration. If decreasing the duration doesn’t produce the correct amount of liquid, change to a needle with a smaller hole.
    Contrastly, if too little liquid is released, increase the duration. If this does not produce the desire amount of liquid, use the fine forceps to gently create a larger hole.

    10) Once the proper volume has been established, inject the embryos at the 1-2 cell stage in the yolk, directly below the cells. Only inject each embryo 1 time, as it is important to maintain a consistent volume of microinjection.

    11) Once all the eggs are injected, turn off the air supply and Picospritzer.
    For Chd-MO injections:
    12) Allow zebrafish to develop for 28 hours.
    13) Anesthetize fish by adding 1X tricaine solution so it fills about half the petri dish.
    14) Place anesthetized fish in small glass vials.
    15) Fix the fish under the hood with BT Fix, making sure that all the E3/tricaine solution is out of the vial first.
    NOTE: BT Fix can be hazardous, so make sure to handle with care under the hood while wearing gloves.
    16) Place fish in BT Fix on the “belly dancer” for 3 hours to allow for complete fixation.
    17) Rinse the fish in 1X PBS 3x for 5 minutes. Dispose of the fix in the bottles under the hood labeled “Fix Waste.”
    NOTE: Fish can be stored in vials filled with PBS in the fix fridge until they are ready to be imaged.

     

    Post-Microinjection

     

     

    Imaging (adapted from Jeffrey Roth’s protocol on the Lom Lab Protocols)

    NOTE: Try to image as soon as possible after fish have been fixed to maintain the integrity of the embryos.
    1) Place fish in a petri dish filled with PBS.
    2) Using forceps, align fish so they are all facing one direction. Avoiding scraping the bottom of the petri dish, as this can mess up the background of the picture.
    3) Place the fish under the stereoscopic microscope located across from the confocal microscope.
    4) Turn on the external light source, microscope (switch on back right), and the Nikon camera. Once the Nikon camera light is green, open NIS-Elements on the computer.

    5) Switch port selector on the left hand side of the microscope from “bino” to “photo.”

    6) Position the specimen you want to image so it is centered on the computer screen. Turn the Zoom knob to 2x so the full image is on the screen.

    7) Adjust the light source as well as microscope light (knob next to the on/off switch) to properly illuminate fish. You can also change the Gain and Exposure to help illuminate the fish. A longer exposure and a higher gain brighten the image.

    NOTE: I placed the external light source arms, at the highest intensity, on each side of the dish. I used an exposure of 1/3 s and a gain of 1.40x. I also turned the microscope light on medium intensity (about half-way).

    8) Use the wheels on the side of the scope to properly focus (large knob = coarse focus, small knob = fine focus).

                NOTE: At the bottom of the NIS-Element window, there is a black bar with an associated number that will move with the focus. The best focus occurs when the bar is farthest left and the highest number appears.

    9) Click “Auto White” to create a clear background.

    10) Once the image is properly illuminated and focused, go to the Camera box and click “Capture.”

    11) File --> Save as “JPG”
    Adjusting the Image
    1) Open the desired image in NIS-Elements.
                NOTE: Camera and microscope must be turned on.
    2) Image --> Adjust Image --> Compensate Background Adaptively --> Degree = 0.01 --> Background = Light (no autocontrast)
                This will create a white background.
    Adding Scale Bars
    1) Take image of the micrometer (found in the shelves next to Dr. Round’s area of the lab). Make sure the zoom on the microscope is the same as the fish images (should be 2x). Also, make sure the resolution in NIS-Elements is the same as the fish images [I used Fast (Focus) = 640x480 normal; Quality (Capture) = 1280x960 normal]
    2) Save image as a JPEG
    3) Open this micrometer JPEG in Adobe Photoshop
    4) On Toolbar, click Select --> All
    5) Edit à Copy
    6) Open fish image (JPEG) to which you want to add scale bars in Adobe Photoshop.
    7) From Toolbar, click Layer --> New --> Layer…

    8) Paste the picture of the micrometer onto this new layer. It should completely cover the fish image.

    9) From the Draw Toolbar on the left, select the rectangle shape. Using the micrometer as your point of reference, draw a rectangle as long as you want your scale bar to be (if doing a whole fish, should be 0.5 mm).

    10) From the Draw Toolbar, click to "T" to create a text box. Add a text box under the recently drawn rectangle (scale bar) and write the length of the scale bar (i.e. 0.5 mm). Make sure this is centered under the bar.

    11) In the box on the right hand side, UNSELECT “Layer 1” so the micrometer disappears.

    12) Click the arrow head button on the Draw Toolbar.
    13) Move the scale bar (with label) directly below the bottom right part of the fish.
    14) Save the image as a JPEG and PSD (in case you need to make any changes).

    • CommentAuthormaangel
    • CommentTimeMay 15th 2012
     
    For protocol with images, go to Public/louise/biology/lom/student research results/ANGEL 13 (Slitrks)/Proposal, Abstracts, and Protocol