Laser Capture Micro-dissection (LCM) of Neonatal Mouse Forebrain for RNA Isolation

Precise and reproducible isolation of desired cell types or layers from heterogeneous tissues is crucial to analyze specific gene profiles and molecular interactions in vivo. Forebrain is the core site of higher functions, like cognition and memory consolidation. It is composed of heterogeneous and distinct cell types, interconnected to form functional neural circuits. Any alteration in the development or function often leads to brain disorders with profound consequences. Thus, precise molecular understanding of forebrain development in normal and diseased scenarios is important. For quantitative studies, most traditional analytical methods require pooling of large cell populations, that results in loss of in vivo tissue integrity and of spatial, molecular and cellular resolution. Laser capture microdissection (LCM) is a fast and extremely precise method of obtaining uncontaminated, homogeneous sets of specific cell types and layers. Our current procedure involves cryo-sectioning and laser microdissection of fresh-frozen mouse forebrains, that are genetically modified and treated with small-molecule therapeutics. Using LCM, specific regions of interest, such as neural layers, cells from adjacent yet distinct subregions within a tissue layer, are obtained under RNase-free conditions. These small cellular cohorts are further used for downstream, high-throughput genomic or transcriptomic assays. Here, we have introduced break-points at multiple stages throughout our protocol. This makes our method simpler and more user-friendly to follow, without compromising on the quality. The current protocol can easily be adapted for different brain regions, as well as for other model organisms/human tissue.


Background
To understand the molecular mechanisms underlying normal and diseased neural processes, it is critical to analyze the interactions in vivo, within a specific tissue microenvironment. It is quite challenging to precisely access different areas of the developing brain, especially in deep areas like hippocampus. The level of complication increases further due to the existence of cellular heterogeneity even within a single neural layer. Traditional quantitative methods like microarray etc., using gross tissue lysates or various primary and organotypic culture systems, rely heavily on pooling gene expression data from thousands of cells and reporting a population-wide average (Levsky and Singer, 2003). Thus, these procedures fail to capture subtle differences within distinct cell subsets as well as to maintain in vivo tissue/ cell integrity. Moreover, there remains a high risk of tissue contamination from neighboring areas during sample collection. Laser-capture microdissection (LCM) combines the benefits of light microscopy and precision of laser beam to procure homogeneous subpopulations of cells from heterogeneous tissue under direct microscopic visualization (Emmert-Buck et al., 1996). The laser cutting is usually less than 1 μm wide. This allows target cells to remain unaffected by the laser beam. LCM technology can selectively and routinely harvest regions of interest, ranging from specific tissue layers to single cells and chromosomes (https:// www.leica-microsystems.com/products/light-microscopes/details/product/leica-lmd7/). This enables the user to obtain reproducible, targeted starting material for biologically relevant data analyses.
Data using the described LCM protocol was published in (Roy et al., 2019). In this study, we found that activating mutations in the phosphoinositide 3-kinase (PI3K) pathway results in focal disruption of cell-cell adhesion, proliferation and apical tissue integrity, leading to cortical gyrification (in both neocortex and hippocampus) and developmental hydrocephalus in mice. Conventional quantitative analytical methods were not precise enough to distinguish cellular/molecular heterogeneity within a neural layer, between adjacent gyral and sulcal zones in the mutant mouse brain. Hence, we used LCM to specifically dissect selective portions of the developing ventricular zone from P0 control and mutant littermates, followed by RNA sequencing. This data was then analyzed to study the effect of the PI3K mutation on neural progenitors. Further, we performed the same experiment after administering a small-molecule drug in utero, to compare the effect of drug treatment on the gene expression profile between controls and mutants.
Here, we present detailed step-wise instructions including experimental 'do's and don'ts', and also illustrate key components and steps using photographs, to aid new researchers in setting up this useful procedure. We have introduced a number of stopping steps throughout the protocol. This makes our method simple and more user-friendly, without making any compromise on the quality, quantity and integrity of the end product (RNA, DNA etc.). This method can be used with fresh-frozen and fixed tissues.
Some other applications of this protocol are as follows:

1.
Absolute identification of possible mosaic mutations with cells of interest, in mouse models or human patients.

a.
Male and female mice are mated, and embryos are staged considering the morning a vaginal plug is detected as embryonic day (E)0.5. In utero drug treatment can be done using this staging. Specifically, the types of mating used in (Roy et al., 2019) are described in detail in the article's Methods section.

b.
The day the pups are born is considered as P0. For FVB strain, the gestation period is E18-E19.

c.
No alterations to the protocol are required if different mouse strains are used.

39.
Nissl stain for RNA research (50 ml) (see Recipes) Note: If the experiment requires comparison of mouse tissue between control and genetically modified mutant models and/or ±drug treatment, reagents to perform mouse tissue genotyping are required as briefly mentioned in (Roy et al., 2015 and2019

21.
UV transilluminator (one used here is from Fotodyne, catalog number: FD33500; should preferably be used only RNase-free and without any chemical contamination)

22.
Rocking shaker (Reliable Scientific Inc., model 55), for solution preparation and staining of slides.

Procedure
Note: Prior to starting laser microdissection, the user should be well trained in cryosectioning and have sufficient knowledge of microscopy and histopathology to sample the correct tissue or cells. To minimize quality damage, the whole procedure of tissue handling should be carried out quickly, yet carefully.

A. Preparation of solutions and set-up for brain dissection
Notes:

a.
Wear sterile disposable gloves and change frequently at every step, from preparing reagents till RNA analysis etc.

b.
Use clean, RNase-free instruments. Before handling the next specimen, spray the instruments with RNaseZap, rinse with RNase-free water and wipe with clean Kimwipes.

c.
Use RNase-free or Nuclease-free solutions, glassware and plastic ware.

d.
Use RNaseZap or similar product to clean the equipment or the surface of the working area.

e.
Freeze tissues immediately after surgery to obtain high-quality RNA. This minimizes gradual RNA degradation by ubiquitous RNases or heat.

1.
Prepare working solutions, as mentioned in the Recipes section

2.
Label the cryomolds at room temperature using the Fisherbrand permanent marker, prior to the tissue dissection experiment, preferably on the side-wall opposite to the notched side (for easier orientation)

3.
Autoclave the dissection instruments and spatula for sterilization, prior to the experiment.
Note: The dissection tools should have never seen any fixative etc.; wipe them with RNaseZap, then RNase-free DNase-free water and dry with clean Kimwipes before starting the experiment.

4.
Wipe the tissue dissection area thoroughly with 100% RNaseZAP solution, let the area dry before placing the laboratory mat for dissection.

5.
Assemble all reagents and equipment before tissue dissection, as shown in Figure  1.

6.
Crush some dry-ice using the mallet to generate a soft powdered area for future placement of the embedding mold ( Figure 2A).

7.
Pour OCT slowly and carefully along one edge of the sterile embedding mold till it is half-filled ( Figures 2B and 2F).
Note: Avoid air bubbles in the OCT while pouring. Air bubbles disrupt the tissue integrity.

1.
Euthanize neonatal mouse pups of the desired age (P0-P6) using procedures permitted by your Institutional guidelines. In Roy et al. (2019), as per the approved animal protocol (see Ethics statement), euthanasia was performed by inducing hypothermia anesthesia, followed by decapitation. For this procedure, place the newborn pups on the lid of a Petri dish and set on top of crushed ice for 2-3 min, leading to their unconsciousness. Confirm anesthesia in pups by lack of response to firm toe pinch, once they stop moving. Decapitate using sharp scissors.

2.
Transfer the heads to a sterile 100 mm Petri dish containing sterile 1x PBS.

3.
At this stage, if genotyping of the pups is required, take a tiny piece of the tail (≤ 2 mm) of each pup and place them in labeled 1.7 ml microcentrifuge tubes for further processing.

4.
Dispose of the carcasses using labeled biohazard bags.

5.
Under a dissection microscope, dissect out the brains using fine and coarse forceps ( Figure 2C). Dispose of the remaining carcasses using the biohazard bags.
Note: If the Petri dish becomes murky with blood, you can transfer the brains at this stage to another Petri dish containing sterile 1x PBS.

6.
After removing the brain, sever the hindbrain and midbrain, retaining only the forebrain including the olfactory bulbs ( Figures 2D-2E). The presence of the olfactory bulb helps to position the telencephalic hemisphere in the correct orientation in the embedding molds.
C. RNase-free brain mounting

1.
With the help of the spatula and coarse forceps (if needed), take the brain out of the buffer and transfer to the pre-labeled embedding mold half-filled with OCT ( Figures 2F-2G).
Note: Try to remove liquid from the brain surface as much as possible without damaging the brain (you may carefully use a finely rolled Kimwipe if needed), before immersing it in the OCT.

2.
Orient the mouse forebrain such that it is parallel to or on the floor of the embedding mold.

3.
For coronal sectioning (perpendicular to the antero-posterior axis of the forebrain), slowly orient the anterior side of the brain (olfactory bulb side) directly perpendicular to the notched side of the mold. Avoid air bubbles in the process ( Figures 2G and 2K).

4.
Once the brain is properly oriented, slowly pour OCT along one edge to fill the mold (or at least enough to cover the entire brain). Take precaution that the brain orientation remains intact and air bubbles do not develop in between OCT layers.

5.
Carefully place the mold within the powdered dry ice and support the mold from the sides by pieces of dry ice ( Figures 2H-2I).

Ethics
Animal experimentation: All animal experimentation was conducted in accordance with the guidelines laid down by the Institutional Animal Care and Use Committees (IACUC) of Seattle Children's Research Institute, Seattle, WA, USA (protocol ID: IACUC00006; last day of triennial approval period: 11 th July 2021). Once the mold, with brain embedded, becomes completely opaque ( Figure 2J), transfer it to a new, clean freezer box and immediately store at −80 °C freezer.

2.
Incubate PEN membrane slides for 30 min under UV light (on the UV transilluminator) to enhance the binding of the tissue to the membrane slide.

3.
Get a bucket of dry ice for tissue/embedded mold transportation.

b.
Nissl stain (Recipe 5, prepare at least 3 days before use and store at 4 °C). Aliquot the amount needed for current use, by roughly estimating the total number of slides to be stained (10-15 drops of the solution is adequate to rapid stain each slide).

5.
Set temperature of cryostat in the range of −15 °C to −18 °C

6.
Label PEN slides with brain code/genotype using a pencil.

1.
Wear new sterile disposable RNase-free gloves. Change gloves frequently between steps to maintain the RNase-free status.

2.
Transfer an embedded brain mold, placed securely on dry ice, from −80 °C freezer to the cryostat.

3.
Use new sterile, single-edge blade to tear the plastic embedding mold inside the cryostat. Trim the OCT-embedded block as needed.

4.
For coronal staining, attach the block on the cryostat chuck (specimen clamp) such that the anterior side of the brain (notched side of mold, Figure 2K) faces perpendicular to the microtome blade inside the cryostat.

5.
Place embedded brain inside the cryostat at least 20 min prior to sectioning, in order to allow it to come to the cryostat temperature.

6.
Use a new sterile microtome blade for each experiment ( Figure 3A).

7.
Slice a few sections from the surface of the mold to avoid any sort of contamination, prior to the collection of actual sections to be mounted on slides.

8.
Cut 8-10 μm sections and mount on regular charged plus slides (on the positively-charged coated side). To identify your region of interest, rapid stain with Nissl staining solution or other dye for about 1 min, swivel gently, and observe slides under stereo microscope.
Note: Staining procedure is explained in Step E14b.

9.
Once you start getting sections of your region of interest, change the slicing thickness to 16-18 μm and mount on the PEN membrane slides. You can mount multiple sections on one slide. For example, ~12-14 sections were mounted on each PEN slide in Roy et al. (2019).
Note: Mount the sections within the extent on the PEN-coated membrane ( Figure  3B). Please take care that the coating does not get any scratches or leave the RNase-free area.

10.
Optimally, place the PEN slides on the desiccant during the section collection.

11.
Clean the blade and cryo-sectioning stage with brush or Kimwipe in between sections.

12.
Collect one section of every 5-10 sections and mount on the regular charged slide for histologic staining.

13.
Air-dry the PEN slides at room temperature for about 2 min on desiccant. Make sure the collected sections are completely dry before staining.

14.
After air-drying, PEN slides with collected sections can be:

a.
Stored in plastic slide mailer in a vacuumed bag, containing desiccant, at −80 °C, for a few months or longer, until user is ready to perform LCM.

b.
Immediately transferred to 100% ethanol at room temperature for dehydration, followed by Nissl staining.

i.
Stain slides one at a time using 50 ml tube or simultaneously using a big container.

ii.
Immerse the slides in fresh 100% ethanol, 5 min x 4 times, and place on the rocking shaker to remove any OCT attached.

iv.
Place drops of Nissl stain (Recipe 5) directly onto the sections, using plastic transfer pipette, and incubate for 1 min. Swivel the slides gently. On average, 10-15 drops of the solution are adequate per slide.

v.
Quickly rinse the slides with fresh 100% ethanol, 5 s x 3 times.

vi.
Finally, immerse the slides in fresh 100% ethanol for 1 min, to allow the stain to bind onto the sections.

vii.
Observe sections under a brightfield dissecting microscope.
viii. Allow the slides to stand for at least 30 min in order to make them completely dry before storage or use.
Note: Optimally, finish staining of sections strictly within 10 min in total.

15.
Store completely dried PEN slides with mounted sections at −80 °C, either before or after Nissl staining. Neither procedure will negatively affect the RNA quality. Keep the slides at −80 °C until you are ready to start LCM.

b.
Copy and paste each image as a separate slide in Microsoft PowerPoint.

c.
Carefully trace the region of interest, using PowerPoint tools, as shown in Figure 3D.

d.
Using these representative tracings, successfully perform LCM on the related PEN slides having the respective serial sections.

5.
If not already done, stain sections on the PEN slides with Nissl stain, as described in the previous section, just prior to microdissection.

6.
Dry the slides post-staining, in the desiccator.

7.
Place PEN slides upside down in the stack-slide holder.

8.
Create a sample overview under 1.25x lens, using LED/halogen illumination.

9.
Mount 0.5 ml RNase-free, DNase-free collection tubes in the LMD tube holder. Add 20-40 μl of a mixture of RLT buffer and 2-mercaptoethanol to the cap of each tube (10 μl 2-mercaptoethanol in 1 ml RLT buffer from Qiagen RNeasy Plus Micro Kit).

11.
Adjust software parameters to get suitable cutting including more accurate and less damage of tissue. Software parameters used in Roy et al. (2019): Draw shapes using "close line". Use the "Move + Cut" mode for cutting respected tissue zones/cells. Use "Fine" X-Y-Z precision mode.

12.
Guide the laser beam with mouse or touchscreen over your specific region of interest. The dissected tissue drops into the caps of collection tube by force of gravity without contamination. In Roy et al. (2019), ventricular zone of P0 hippocampal CA1 field (CA1 vz) was collected from each coronal hemisphere ( Figure 4).
Note: Please load a new collection tube every 2 h if longer cutting time needed.

13.
Once all sections of the first slide are laser-cut, move to the next slide and repeat Steps F9-F12. You can store the used slides for future inspection and documentation.
Note: Depending on the size and density of the laser-dissected samples, you can decide whether to pool together samples obtained from each brain for RNA isolation and further processing. In Roy et al. (2019), our region of interest (ventricular zone of P0 mouse CA1 region) was both low in volume and in density. Hence, total RNA was isolated from LCM-enriched samples pooled across ~6 slides per genotype.

14.
After microdissection, carefully retrieve the collect tubes, close the cap upside down and vortex for 1 min.

15.
Give the tubes a brief spin in the centrifuge in order to collect sample at the bottom of the tube. Store tubes at −80 °C for further analysis.

c.
Shake the solution for a few hours and store at 4 °C, sealed air-tight and dark for up to 6 months. This solution should be prepared at least 3 days prior to usage. Aliquot as required and discard the used solution. Reagents and equipment needed for mouse brain dissection are labeled. 1x PBS-Phosphate buffer saline, working solution.  A. Crushed dry ice prepared for freezing the embedding mold. B. OCT poured along an edge of the embedding mold to avoid air bubbles. C-E. Dissection of neonatal mouse forebrain, in sterile 1x PBS. F-G. Forebrain carefully transferred to and then oriented in the embedding mold filled with OCT. H-J. Mold with OCT-covered oriented mouse forebrain placed in dry ice, until OCT freezes to opaque form. K. Schematic of how forebrain was oriented, to be used for coronal sectioning (Roy et al., 2019). A. New sterile microtome blades used for brain sectioning. B. PEN slides: mount sections onto the inner film, which finally gets laser-cut. C. Home-made desiccator using Drierite to dry slides prior to LCM. D. Representative image of a Nissl-stained coronal hemi-section of P0 mouse forebrain, used in microdissection and data analysis (Roy et al., 2019), showing how region of interest is traced. In this context, ventricular zone of P0 mouse hippocampal CA1 region (CA1 vz) was traced.  Different steps-tissue sectioning, laser-capture microdissection, downstream applications. Some images are obtained from Roy et al. (2019), where we compared the genetic profiles of the hippocampal CA1 ventricular zone (CA1 vz) between P0 control and activating Pik3ca mutant ± drug, using RNA sequencing. Roy et al. Page 19 Bio Protoc. Author manuscript; available in PMC 2020 July 05.

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