Quantitative Analysis of Protein Expression to Study Lineage Specification in Mouse Preimplantation Embryos

Ethics statement: All animal work, including husbandry, breeding and sacrifice was approved by Memorial Sloan Kettering Cancer Center's Institutional Animal Care and Use Committee (IACUC), protocol #03-12-017.

1. Embryo Collection

Note: All animal work must have been approved by institutional and local authorities and conform to local and institutional rules.

1. Mate a virgin female mouse with a fertile stud male of the desired genotypes.
   Note: If setting up natural matings, selecting females in the estrus phase of the estral cycle increases the chances of copulation on the desired date. If inducing superovulation, please refer to the protocols described in ²⁰.
2. Check the presence of a vaginal plug in the morning using a blunt probe. Ideally, do this before noon (12:00 pm), as copulation plugs are lost throughout the day. Consider noon of detection of the vaginal plug embryonic day (E)0.5.
3. On the desired day and time of embryonic development, warm up M2 or flushing holding medium (FHM) to RT or, preferably, 37 ºC. Note: Either M2 or FHM can be used for flushing and handling embryos. When not in use, store these media at 4 ºC. Estimate the use of ~2 ml of medium for each uterus.
4. Thaw frozen 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) or prepare some fresh solution. Estimate 500 µl of 4% PFA solution per well of a 4-well plate. Fix one litter on each well of a 4-well plate. Note: Alternatively, a 96-well plate can be used to fix and store embryos. If using a 96-well plate, use 100 µl of PFA solution per well.
5. Pull a glass Pasteur pipette using an open flame to draw a capillary end at the tip.
6. Sacrifice female by cervical dislocation or CO₂ inhalation, or as required by local and institutional regulations. It is often desirable to confirm euthanasia of the animal by cervical dislocation.
7. Spray belly of the animal with 70% ethanol to minimize hair shedding and perform an abdominal incision, first through the skin and subsequently through the body wall (peritoneum) to expose the viscera.
   Note: The following steps describe the procedure to collect blastocysts from the uterus of a pregnant female mouse at any stage between E3.25 and E4.5 (Figure 1A). For protocols on collection of preimplantation embryos earlier than E3.25, refer to ²⁰.
8. Locate the uterus and remove from the animal. Holding the cervical end of the uterus with a pair of forceps, cut through the cervix and pull up gently to stretch both uterine horns.
9. Trim fat from the uterus, taking care not to pierce the uterine wall. This can be done easily at the time of removal, while still attached to the body of the animal, as the uterus can be stretched out. Alternatively, it can be done later, under a dissection microscope, in RT PBS. For a more detailed protocol on dissection of the uterus, see Protocols 5 & 8 ²⁰
10. Cut above the oviducts, below the ovaries, to release the entire uterus, place on a petri dish and cover with PBS.
    Note: This step also allows the washing of excess blood or other debris from the uterus, which facilitates subsequent blastocyst harvesting.
11. Separate both uterine horns by cutting through the proximal end, on both sides of the cervix and discard the cervix. Separate each oviduct from the uterus by cutting below the isthmus that connects them (Figure 1A). Transfer both horns to a drop of manipulation medium (either M2 or FHM) for embryo collection and manipulation.
12. Under a dissection microscope, flush blastocysts out of the uterus and into the medium by forcing 0.5 – 1 ml of M2 or FHM through each uterine horn from the cervical opening. Use a 1 ml syringe with a hypodermal needle. Note: The needle can be blunted using a filing stone to avoid tearing the uteri, although this is not critical. A detailed diagram for uterine flushing can be found in²¹.
13. Using the dissection microscope, locate the blastocysts, which will be scattered throughout the drop of medium. Allow up to 1 – 2 min for blastocysts to sink to the bottom of the dish after flushing. Using the mouth-controlled, glass Pasteur pipette with a capillary end, collect all blastocysts and transfer to a fresh drop of media.
14. Rinse blastocysts by moving them through 2 – 3 drops of fresh M2 or FHM. Move embryos in groups of 5 – 10 blastocysts at a time.
    Note: Moving larger groups at any one time will speed up the process, but it will also increase the chances of accidentally losing many embryos. If untrained in the use of mouth-controlled pipettes, practicing embryo manipulation and transfer should be considered before beginning the experiment.
15. Remove the Zona Pellucida (ZP) from unhatched blastocysts (generally <E4.0) by briefly washing them in acidic Tyrode's solution. As soon as the ZP is no longer visible, transfer blastocysts back to manipulation media. Only keep embryos in acid Tyrode's for as long as strictly necessary, in order to prevent damage to the cells.
    1. Take care when handling denuded blastocysts, as they adhere very easily to glass and plastic surfaces. Manipulation medium contain 4 mg/ml of bovine serum albumin (BSA) to prevent attachment, but acid Tyrode's or PBS do not, therefore, to reduce the risk of damage or loss of the embryos do not pick up more than 2 – 5 denuded blastocysts at a time when manipulating them in BSA-free or serum-free PBS or acid Tyrode's.
       Note: Alternatively, coat the glass pipette with 1% BSA before use to reduce adherence of the embryos to the glass surface.
16. Coat the bottom of each well of a 4-well tissue culture dish with a thin layer of 1% agar, 0.9% NaCl to prevent embryos from adhering to the plastic. Alternatively, add 4 mg/ml of BSA to the PBS (PBS-BSA).
17. Fill one well of the coated 4-well tissue culture dish with 500 µl of RT PBS (or PBS-BSA) and another well with 500 µl of 4% PFA in PBS.
18. Rinse blastocysts in RT PBS and fix in 4% PFA in PBS for 10 min at RT. Transfer to PBS (or PBS-BSA) after fixation.
19. If embryos are not going to be processed immediately, cover the PBS containing the embryos with a layer of mineral oil to prevent evaporation (leading to desiccation of the sample) and store the plate at 4 ºC.
    Note: Different fixation methods and times can be used depending on the experiment, the epitopes of interest and the antibodies to be used. Whichever the method of choice, be consistent throughout equivalent experiments to facilitate subsequent comparison of the staining results.
    Note: Pause point: Embryos can be stored at 4 ºC in PBS for up to several weeks before proceeding to the next step. Ensure PBS is sterile and/or add 100 U/ml of Penicillin + 100 µg/ml Streptomycin (Pen-Strep) to prevent bacterial contamination (especially when using PBS-BSA) if anticipating prolonged storage.

2. Immunofluorescence

1. Perform immunofluorescence using any protocol that has previously been tested and proven to be robust for whole-mount immunofluorescence. Note: We recommend those described in ²² and ¹¹. In the present article the former will be described. Whichever the method of choice, be consistent throughout equivalent experiments to facilitate comparison of the staining results.
2. Use a flexible, polyvinyl, clear, U-bottom 96-well plate to carry out the sequential steps of the immunofluorescence protocol. Fill each well with 50 – 100 µl of solution and move embryos from one solution to another using a L-shaped mouth pipette. Note This approach minimizes the use of reagents and facilitates tracking steps as well as the parallel staining of several experimental groups.
   1. Optional : Coat the bottom of the wells with a thin layer of 1% agar, 0.9% NaCl to prevent the adhesion of embryos to the plastic. Do not add BSA to immunofluorescence solutions.
3. Prepare PBX (0.1% Triton X-100 in PBS). If anticipating the performance of several immunofluorescence experiments, prepare 50 ml of PBX and store at 4 ºC between experiments.
4. Prepare permeabilization solution (0.5% Triton X-100, 100mM glycine in PBS). Make 1 ml (or whichever amount necessary) of fresh permeabilization solution for each experiment.
5. Prepare blocking solution (2% horse serum (HS) or 20% fetal bovine serum (FBS) with Pen-Strep in PBS). Prepare 10 ml and store at 4 ºC between experiments.
   1. No differences have been observed between either type of serum, however, be consistent in the choice of blocking solution used for equivalent experiments.
6. Rinse fixed embryos for 5 min at RT in PBX (~100 µl).
7. Permeabilize embryos for 5 min at RT in permeabilization solution (~100 µl).
8. Rinse embryos for 5 min at RT in PBX (~100 µl).
9. Block embryos for 30 min to 1 hr at RT in ~100 µl of blocking solution. Cover the solution with a layer of mineral oil to prevent evaporation or keep in a humidified chamber.
   Note: Embryos can be blocked for up to O/N periods at 4 ºC without noticeable improvement or detriment when compared to 30 min blocking steps.
10. Incubate embryos O/N at 4 ºC in primary antibody/antibodies diluted in blocking solution (~100 µl). Cover the solution with a layer of mineral oil to prevent evaporation or keep in a humidified chamber.
    1. Determine optimal dilution for each primary antibody beforehand. See discussion and Materials section for tested dilutions of a number of antibodies.
11. After O/N incubation, rinse embryos 3x for 5 min each at RT in PBX (~100 µl).
12. Block embryos for 30 min to 1 hr at RT in ~100 µl of blocking solution. Cover the solution with a layer of mineral oil to prevent evaporation or keep in a humidified chamber.
13. Incubate embryos for 1 to 2 hr at 4 ºC in secondary antibody/antibodies diluted at 4 µg/ml in ~100 µl of blocking solution. Cover the solution with a layer of mineral oil to prevent evaporation or keep in a humidified chamber.
14. Rinse embryos 2x for 5 min each at RT in PBX.
15. Move embryos to preferred DNA stain. If using Hoechst 33342, dilute to 5 µg/ml in PBS. Cover the solution with a layer of mineral oil to prevent evaporation or keep in a humidified chamber.
    Note: Pause point: If embryos are not being imaged immediately, they can be kept for up to several weeks in nuclear staining solution. In this case, ensure PBS is sterile and/or add Pen-Strep or Sodium Azide to prevent bacterial growth and contamination.

3. Confocal Imaging

Note: Individual confocal microscope configurations will require specific acquisition parameters to be adjusted for the system and software in place. However, the following section provides a set of general rules to follow that should be applicable to any given installation.

1. Using a fine mouth pipette, make microdrops of PBS or nuclear stain solution on the glass surface of a 35mm glass-bottom dish and cover with mineral oil. As an alternative, use a regular coverslip with silicone separators to avoid damaging the embryos and cover with a glass microscope slide. Note: When using a regular coverslip, embryos cannot be subsequently re-positioned or recovered for genotyping, therefore we strongly recommend the use of glass-bottom dishes.
2. Place embryos in the microdrops and arrange in a consistent manner, preferably with the ICM-cavity axis parallel to the glass surface. Set up dish on the microscope holder. Note: Images obtained on laser scanning confocal microscopes suffer from a Z axis-associated loss of fluorescence intensity. Therefore, consistency in the arrangement is important for comparing intensities between equivalent regions on different embryos.
3. Set up reference parameters using wild type embryos that have not been subject to any sort of treatment that may alter gene expression.
   1. Acquire images using a high magnification objective (i.e., 40X or higher) in order to obtain data that will facilitate accurate segmentation using the MINS tool ⁴.
      Note: Using digital zoom on a 20X objective will not yield images of adequate resolution for segmentation using MINS. One should also note the working distance (WD) of the objective, as it should be sufficient to allow acquisition of a Z-stack that spans an entire blastocyst, thus a long WD (~130 – 150 µm) objective is usually necessary. The images shown in Figures 2 – 4 were acquired using a 40X/NA 1.30 oil immersion objective with a working distance of 210 µm.
   2. Use the lowest laser output that provides a strong signal-to-noise ratio without bleaching the fluorophores.
   3. Adjust the gain and offset to obtain the widest dynamic range without overexposing the sample. Exposing the images to cover most of, or span the entire, gray scale range will facilitate the detection of small differences in intensity between images.
   4. Use a small (1 µm) Z-step, since Z axis resolution is critical for accurate segmentation with MINS.
      Note: XY dimensions do not affect the nuclear segmentation process, only the image file size. Generally, 512 x 512 pixels is a sufficient XY resolution for publication-quality images without compromising hard-drive space.  
   5. Critical step: Keep imaging parameters consistent within and across imaging sessions of the same experiment so as to be able to compare samples.
4. Imaging batches of embryos:
   1. Image coherent groups (litters, experiments) in a single session whenever possible.
   2. Keep parameters consistent within and across imaging sessions for replicates of the same experiment.
   3. Image embryos throughout (whole Z axis) to capture every cell.
      Note: If desired, embryo genotyping may be done after imaging as described in ²³. The need for nested PCR has to be determined empirically and will depend on the locus to be analyzed and the primers used.
5. Make sure to be able to match embryos to images retrospectively.

4. Image Analysis and Data Pre-processing

1. Download and install MINS ⁴ from http://katlab-tools.org (MATLAB license required).
2. Perform image segmentation:
   1. Follow the graphic user interface (GUI) of MINS to load a confocal image or stack from the hard drive, detect the nuclei and segment the image. Load the entire Z-stack of raw data generated by the microscope (.lsm, .lif, .oif, etc.). See Table 2 for details on each step and instructions found on  http://katlab-tools.org and⁴. Once completed, view the outcome of each step by clicking the corresponding 'View' button. A yellow tag above the button indicates operation in process, a green tag indicates operation completed.
      Note: MINS currently does not support .czi files, .tiff sequences or multi-position files – each Z-stack needs to be an independent file.
   2. After satisfactory segmentation, save the pipeline created so it can be used directly for other embryos or litters.
3. Batch-Mode segmentation:
   Note: Single files can be segmented individually. However, given that embryos within a litter or experiment are generally of comparable stage, MINS can apply the segmentation pipeline created for a single image to a group of them without supervision.
   1. From the Batch-Mode Run menu, click 'Add files' and load all files to be processed at once. Note: Files from different sources can be added to the segmentation queue.
   2. Click 'Start Batch-Mode Run' and allow time for the software to process the files. Note: Segmentation output will be saved in the same directory where the original files where located.
4. Segmentation assessment and manual correction
   Note: MINS segments images with very high accuracy (>85%), however, the quality of the staining and imaging, as well as the stage of the embryo, may affect MINS performance. Generally, the higher the nuclear density within the embryo, the more likely segmentation errors will take place (Figures 2D, 3). Thus, segmentation accuracy should be evaluated for each sample and manually corrected if needed. Two types of errors typically occur: over-segmentation and under-segmentation.
   1. Resolving over-segmentation:
      1. Identify false positives – apoptotic vesicles (Figure 3A a, b, d, asterisks) or other elements that are not intact nuclei, but which may have been identified as such by MINS.
         Note: Generally such false positives have a smaller size than real nuclei and can, in the majority of cases, be easily identified and sorted on the spreadsheet. However, visual examination is strongly recommended, as oftentimes nuclei may be only partially segmented, resulting in a smaller, but valid segmented volume (Figure 3A b, arrowhead). 
      2. Delete the corresponding records for false positives from the *statistics.csv file. Preserve the original *statistics.csv file for future reference and edit only a copy of it.
      3. If a nucleus has been over-segmented and presented as 2 or more nuclei (Figure 3A c, arrow and arrowheads), either merge the records by averaging their intensity level or, if similar, keep one of the records for that cell and discard the rest.
   2. Resolving under-segmentation:
      1. Identify events where MINS has failed to detect the border between two or more nuclei and consequently segmented them as a single nucleus (Figure 3A d, arrow and 3B). Note: This poses a problem for an accurate cell count, but more importantly, for quantifying protein levels, as the levels measured will be the average of the cells that have been erroneously merged, and which may belong to different lineages.
      2. Identify events where MINS has failed to detect a nucleus altogether.
      3. In these instances (4.4.2.1 and 4.4.2.2), measure the average gray levels for each channel in the under- or un-segmented cell(s) using an alternative tool, such as ImageJ (http://imagej.nih.gov/ij/), and replace the erroneous record with the correct levels. Preserve the original *statistics.csv file for future reference and edit only a copy of it. To do this using ImageJ, follow these steps:
         1. Open the relevant multi-channel stack on ImageJ and import the corresponding *overlaid.tiff file generated by MINS as a virtual stack ('File' > 'Import' > 'TIFF virtual stack…').
         2. Find a medial section of the under- or un-segmented nucleus.
         3. Using the freehand selection tool outline the perimeter of the under- or un-segmented nucleus on the DNA stain channel.
         4. Press Crl+M (Cmd+M on a Macintosh) or go to the Analyze menu and select 'Measure'. This action will record the Mean gray value for the area outlined and will display it on a new window. Go to 'Analyze' > 'Set measurements…' and make sure 'Mean gray value' is selected. Select any other parameters to be measured.
         5. Using the same area outlined in step 4.4.2.3.3, repeat the measurement for each of the fluorescence channels of interest. The results will be appended to the previous one on the measurements 'Results' window.
         6. Use the measurements obtained to replace the erroneous records in the *statistics.csv file. If the nucleus had not been segmented, insert a new row, assign it a new Cell ID and introduce the values obtained in ImageJ under the corresponding column for each channel. This cell will not have spatial coordinates (X, Y and Z values). If the nucleus had been undersegmented, duplicate its row, assign different Cell IDs to each new cell and introduce the values obtained in ImageJ under the corresponding column. In this case, both cells will share spatial coordinates.
            Note: If spatial distribution is to be analyzed, we recommend excluding the XYZ coordinates of these cells, as they will overlap. To this end, when creating new records, assign Cell IDs to them that are easily distinguished from the original ones (for instance, non-integer numbers).
   3. Score dividing cells. Note: MINS detects mitotic as well as interphase nuclei, but cannot distinguish between them.
      1. If mitotic nuclei are to be considered separately in the analysis, manually score these and add this information to the data file. To record mitotic cells, add a new variable (column) to the *statistics.csv data file and assign different values to mitotic and interphase nuclei.
5. Data pre-processing.
   Note: Once the spreadsheets have been corrected for over- and under-segmentation they are ready to be analyzed. The analysis process and data presentation will depend on the specific experiment. However, below are some general data transformations we recommend carrying out prior to analysis:
   1. Transform the fluorescence intensity values into their logarithm or square root to reduce the scatter of the data. This also helps visualization, as the raw data tend to concentrate near the axes of the plot (see Figure 4B, C).
   2. Correct the Z-associated attenuation of fluorescence:
      Note: Fluorescence intensity in laser scanning confocal microscopy images decreases the deeper on the Z axis a slice is located (Figure 4E, F).
      1. If one of the epitopes detected in the samples is a ubiquitous and homogeneously expressed housekeeping nuclear marker, normalize the fluorescence intensities of the other epitopes by dividing their values in each cell by the corresponding value of the housekeeping marker.
      2. In the absence of such a reference marker, compensate Z-associated loss of fluorescence in the following manner:
         1. Plot the values of each marker (Y axis of the graph) over the Z position (X axis of the graph) to visualize the decrease in intensity over Z – plot together the values for all control, wild type embryos (Figure 4F).
         2. Fit a linear model (regression curve) to the plot obtained in the previous step (intensity values over Z) for each marker (Figure 4F).
         3. Correct all values for each marker using the following formula:
            log(original value) + Z * slope of the model (Figure 4D, G).
            Note: The specific steps to perform this correction will depend on the analysis software used (R, MATLAB, etc.). If using R, run the following formula to fit a linear model to the data:
            >lm(log(channel)~Z, data = dataframe) 
            where, channel is the fluorescence channel to be corrected, Z is the Z coordinate and dataframe is the table containing all the values for the desired embryo(s) (the *statistics.csv file generated by MINS or a modification of it).
            Note: The output of the above formula will have the following format:
            (Intercept)    Z
            4.76373       -0.01947
            where, the value Z is the slope of the regression line for that dataset, which absolute value should be used to modify the original value with the formula given in 4.5.2.2.3 (see Figure 4G for an example).