레이저 캡처 서 유전자 표정 분석에 대 한 마우스 눈에서 높은 순수 배수 채널의

The overall goal of this laser microdissection procedure is to develop a robust and reproducible method for the isolation of a highly-enriched trabecular meshwork from mouse eyes using laser capture microscopy for subsequent RNA isolation and downstream expression analysis. This method can help answer key questions in the ophthalmology field such as what are the important genes and signaling pathways that, when mis-regulated, can lead to ocular diseases. The main advantage of this technique is that high-quality RNA from the trabecular meshwork of mouse eyes can be reproducibly isolated for gene expression analysis.

Generally, individuals new to this method will struggle because laser microdissection is powerful technique which involves various troubleshooting steps throughout the entire protocol. Begin this procedure with optimal tissue collection for laser microdissection as described in the text protocol. Spray RNase decontamination solution onto the brush, forceps, coupling jars, and mounting chuck.

After wiping thoroughly, rinse the apparatus with nuclease-free water and dry. Wipe down the cryostat with 100%ethanol to avoid cross-contamination. Squeeze a small amount of O.C.T.onto the mounting chuck, then remove the tissue block from the mold and carefully place the block on the mounting chuck.

Once the tissue block on the mounting chuck is frozen solid, insert onto the specimen holder of the cryostat. Next adjust the cryostat to cut eight-micron sections and carefully section through the O.C.T.block to reach the tissue sample. Once the tissue is observed, stop sectioning.

Trim the frozen blocks on all sides using a clean razor blade and leave three to four millimeters of O.C.T.surrounding the tissue to enable handling with the paint brush. Section through the eye, working quickly. Stain every third section by flooding the slide with a quick one-step H&E stain for 60 seconds.

Rinse the section with tap water and air dry. After clearing the section with a clearing agent, mount the section on a charged glass slide with a cover slip, review under the microscope. Continue sectioning until the TM becomes evident in the stained sections.

Once the TM starts to appear, cut two sections and mount them onto a charged glass slide for routine H&E staining to form a map for cutting with laser capture microdissection. After the first H&E map slide, cut and line up six serial eight-micron-thick sections in the cryostat and simultaneously mount them onto the PET membrane slide. Adhere the tissue by briefly sliding a gloved thumb along the back of the membrane.

After mounting, place the PET membrane slide immediately into a slide box on dry ice. Continue to section the eye. After every two PET membrane slides with six sections each, collect two sections on a charged glass slide to create another map slide for H&E staining.

Continue sectioning approximately 100 serial eight-micron-thick sections until the TM is no longer visible. Confirm the lack of TM by quickly staining one section on a charged glass slide. After performing the H&E map slide staining as described in the text protocol, review H&E slides visually using a microscope.

Identify map slides with TM clearly visible and accessible for cutting. Use the PET membrane slides between these map slides for LCM. Remove the slide box containing frozen tissue sections from the minus-80 degrees Celsius freezer and place it on dry ice.

Process the slides one at a time. Fix the sections by placing the slide immediately into cold 75%ethanol containing RNase inhibitors for 30 seconds. Gently dip the slide in nuclease-free water for 10 to 15 seconds to dissolve the O.C.T.without interfering with the tissue sections.

Carefully pipette 300 microliters of 1.5%cresyl violet acetate solution directly onto the sections and incubate for 45 seconds at room temperature. Then wash the sections one time by placing them into a 75%ethanol bath for 30 seconds. Pipette 200 microliters of Eosin Y solution onto the sections and incubate for three to five seconds.

Dehydrate the tissue by placing the slides subsequently in 75%ethanol, 95%ethanol, and finally a 100%ethanol bath for 30 seconds each. Blot the end of the slide with a tissue wipe between every step to remove excess liquid. Let the slide air dry completely under the hood for five minutes.

Once dry, perform LCM immediately thereafter. Avoid the use of xylene for the last clearing step. Xylene treatment will cause the tissue section to inadequately mount to the membrane slide.

Xylene will also reduce the efficiency of the tissue capture by LCM. Place the PET membrane slide with the stained tissue side facing down directly on the top of a new glass slide that has been loaded onto the microscope stage. Make sure the membrane and the glass slide are paired tightly on the microscope stage.

Start LCM by first setting the slide limits. Choose the lowest 4X magnification in the Objective panel, then define the work area of the membrane slide by moving the microscope's XY stage until the upper left corner, where the metal and membrane meet, is visible in the live video feed. At this point, press the Limit 1 button and a red rectangle will appear on the road map.

Next move the XY stage to the opposite corner and press the Limit 2 button. Press the Scan button to create a road map image of the membrane and tissues between the set limits. Double-click near the TM of one of the eye sections within the road map.

The TM can be located near the apex of the iridocorneal angle. Once the TM has been identified, increase the magnification to 10X and manually focus on the TM.Repeat with 20X and 40X objectives. When the TM is in focus with the 40X objective, navigate into a blank area, select the freehand drawing tool, and draw a long line on a blank area of tissue.

Set the laser parameters so that the cut velocity is 34%focus is 58%and power is 70%and then press the Cut button. Make fine laser adjustments to the speed, focus, and power parameters until a clear and fine cutting line is observed. For precise cutting, ensure cutting action follows the drawn line.

Next load the isolation tube lid into the cap holder and open the tube. Attach the cap holder to the cap lift, and make sure the tube is inverted. Also assure that the adhesive cap material protrudes beyond the tip of the cap to ensure that the desired tissue is properly captured.

It is critical to visually ensure that the trabecular meshwork is picked up and attached to the isolation cap. Select the Manual Dissection mode on the software panel. Carefully review that the TM is directly beneath the isolation tube.

Set the White Balance and adjust the Brightness to acquire optimal image quality. To begin cutting, manually adjust the focus and draw a line using the freehand tool. Be sure that the collection cap remains in the up position, not yet touching the membrane.

Draw partial circles near where the TM meets the sclera and press Cut. Once the first cuts are done, position the cap into the down position and readjust the focus, then draw two lines in the open or free space to connect the two partial circles, completing the circle around the TM.Then press Cut and collect tissue from the section. Ensure that the TM was picked up by the microdissection cap.

Position the cap back into the up position and repeat on the remaining sections. Slightly adjust the cap position so that all the isolated samples will fit onto the cap and not overlap. For the next PET membrane, insert a new isolation tube and repeat this section.

To perform lysis of the microdissected TM tissue, carefully pipette 10 microliters of lysis buffer onto the lid of the collection tube, ensuring the lysis buffer completely covers the microdissected TM.Gently close the adhesive cap and incubate the collection tube upside down at room temperature for 10 minutes. After incubation, centrifuge at 800 g for two minutes and place the lysate on dry ice. Store the lysates at minus-80 degrees Celsius for later purification and analysis.

Rapidly degrading RNA can impact the downstream RNA analysis. RNA quality should be measured and quantified using a microfluidics-based platform. Representative results of gene expression analysis of the high-quality total RNA purified from the trabecular meshwork are shown here.

The digital gel for the remaining total RNA from the eye tissue after laser microdissection shows a high RNA integrity number, which is needed for the subsequent downstream RNA analysis. Once quality RNA is extracted from the tissue, downstream analysis such as micro-array, RNA-Seq, and qPCR can be performed. Here, quantitative digital PCR analysis was used to analyze the expression of the trabecular meshwork-associated genes, myocilin and ACTA2 from LCM-collected RNA.

It is illustrated by the heat map generated from data obtained from micro-array analysis that this technique can be used to understand gene expression changes in the trabecular meshwork of wild type mice and knockout mice with an elevated IOP phenotype. Once mastered, this technique can be done in as little as one day per biological animal if done properly. Visual demonstration of this method is essential as the laser microdissection steps are difficult to learn due to the difficulty of the visualization of the cells of interest and the absence of standards for these unique samples.

Relying on the skilled operator is critical. While attempting this procedure, it's important to remember that RNA in small amounts of tissue can be rapidly degraded. Precautions such as decontaminating equipment from RNase, minimizing exposure to air, and keeping tissue cold should be taken.

Though this method can provide insight into the effects of increased intraocular pressure on the trabecular meshwork-specific gene expression, it can also be applied to other systems, such as evaluating changes in gene expression in the retina caused by increased intraocular pressure. Following this procedure, other methods like RNA-Seq or micro-array can be performed in order to answer additional questions like what signaling pathways are altered in the trabecular meshwork of mice with elevated intraocular pressure. After its development, this technique paved the way for researchers to more effectively study gene expression changes in the trabecular meshwork in mouse models.