Combining mass spectrometry and genetic labeling in mice to report TRP channel expression

Transient receptor potential (TRP) ion channels play important roles in fundamental biological processes throughout the body of humans and mice. TRP channel dysfunction manifests in different disease states, therefore, these channels may represent promising therapeutic targets in treating these conditions. Many TRP channels are expressed in several organs suggesting multiple functions and making it challenging to untangle the systemic pathophysiology of TRP dysfunction. Detailed characterization of the expression pattern of the individual TRP channels throughout the organism is thus essential to interpret data such as those derived from systemic phenotyping of global TRP knockout mice. Murine TRP channel reporter strains enable reliable labeling of TRP expression with a fluorescent marker. Here we present an optimized method to visualize primary TRP-expressing cells with single cell resolution throughout the entire organism. In parallel, we methodically combine systemic gene expression profiling with an adjusted mass spectrometry protocol to document acute protein levels in selected organs of interest. The TRP protein expression data are then correlated with the GFP reporter expression data. The combined methodological approach presented here can be adopted to generate expression data for other genes of interest and reporter mice.• We present an optimized method to systemically characterize gene expression in fluorescent reporter mouse strains with a single cell resolution.• We methodically combine systemic gene expression profiling with an adjusted mass spectrometry protocol to document acute protein levels in selected organs of interest in mice.


Preparation of the 2D expression atlas
Step 1: Perfusion of animals and dissection of organs Material • 4% paraformaldehyde (PFA) in PBS • anesthesia according to your country's animal laws • 18% sucrose in PBS • 0.5 M EDTA All animals, regardless of age, are transcardially perfused and all organs are removed. All dissected organs are kept in 4% PFA on ice for three hours. The thorax package is stored in a 50 ml falcon tube, the pituitary and trigeminal ganglia in a 1.5 ml reaction tube, while the rest are stored in 15 ml falcon tubes [1,2] .
1. Fill one syringe with PBS and the other with PFA, place the PFA syringe on ice. 2. Anesthetize animals and wait until reaction to a toe pinch is no longer observed before cutting the chest open and exposing the heart. We use a mix of xylazine (32 mg/kg) and ketamine (200 mg/kg). 3. Make a cut in the right atrium, place the needle in the left ventricle and perfuse the animal with PBS (volume depends on the age, until the liver is clear and no more blood comes out of the atrium) followed by ice-cold 4% PFA (60 -80 ml for adult, 20 -40 ml for juvenile). 4. Remove the skin on top of the salivary gland ( Fig.1 A) and cut out the salivary gland. 5. Cut the trachea close to the tongue ( Fig. 1 B) and lift it up. With a blunt edged scissor cut behind the trachea and remove the thymus, heart and lung together with the trachea as one package ( Fig. 1 C). 6. Remove a piece of liver including the gallbladder ( Fig. 1 D). You only need a small piece of liver including the gallbladder. The rest of the liver can be cut away. 7. Cut the duodenum close to the stomach ( Fig. 1 E) and remove the whole intestinal tract, cut at the end of the colon ( Fig. 1 F). 8. Cut out a piece of duodenum, jejunum, ileum and colon and remove the cecum ( Fig. 2 ). 0.5 mm in length for each intestinal compartment is sufficient. 9. Dissect the stomach and cut away the attached pancreas ( Fig. 3 A). When removing the pancreas, you can leave the spleen attached ( Fig. 3 B). 10. In females, remove the bladder, and subsequently the gonads (uterus with ovaries) ( Fig. 3 C). 11. In males, first expose the testes by pushing them out and cut the vas deferens to remove the testes with epididymis ( Fig. 3 D). Next, dissect the seminal vesicle and cut the bladder out with the prostate attached ( Fig. 3 E). 12. Next, remove the kidney with the adrenal gland attached ( Fig. 3 F). 13. Cut the head off at the neck, make a cut through the skin along the midline and fold the skin to the side. Make three cuts into the skull with bone scissors, one along the midline and two on the side ( Fig. 4 A) and carefully remove the top of the skull with forceps. Remove the bones on top of the olfactory bulb and lift the brain out of the skull cavity by inserting forceps at the back of the brain under the cerebellum, lifting the brain forward, while flipping it towards the nose. The still attached ocular nerves can be gently pulled loose using forceps, to release the brain ( Fig. 4 B). 14. To remove the pituitary gland ( Fig. 4 D), gently loosen the pituitary by pushing against it with forceps. You can pick it up with forceps or wash it into a 1.5 ml reaction tube with PFA and a pipette. You can also dissect the attached ganglia by carefully cutting out the marked area in Fig. 4 D with a scalpel. (Note: this method only works when the animal is fixed, in unfixed animals the pituitary will break). 15. To dissect the tongue, first cut the lower jaw away and then cut the tongue at the epiglottal region to remove it. 16. Next, remove the remaining skin from the head and take out the eyes from their sockets then use fine forceps to remove the hard palate ( Fig. 4 F) and expose the olfactory tissue. 17. To prepare the nose, remove any remaining skull and as many muscles as you can. Leave the teeth in place until after decalcification.
After three hours post fixation in 4% PFA, change the organs into 18% sucrose and keep them at 4 °C until the following day. Do not keep the organs in sucrose longer as it leads to sectioning difficulties. The nose is transferred into 0.5 M EDTA for a 3 to 4 days at 4 °C before it is placed in 18% sucrose (in this case for only 3 hours before freezing, see also below).
Step Step 2.1: Organ freezing We suggest incubation of all organs for three hours at room temperature (RT) in OCT before freezing. For this, you can use standard 24 well plates but it is recommended to place each organ in its own well as this improves sectioning results. Before transferring organs from sucrose to OCT, blot the sucrose away using tissue paper and dissect some organs further (see below, Point one). We recommend the isobutane/dry ice method, described below, over other freezing methods as the tissue block freezes more homogeneously and it affords better control over organ orientation.   1. Before placing in OCT wells, try to dissect as much fat away from the organs as possible as fat will interfere with sectioning. Cut the trachea off the thorax package just above the thymus and put these two pieces separately into OCT wells. Separate the pancreas from the spleen and place each organ into a single well. Make a small cut into the stomach and remove the food before placing in the OCT well and try to pump OCT into the heart and stomach by gently squeezing the organ with feather forceps. 2. Our recommended embedding molds have plastic demarcations marking the front of the tissue block. If you are using different ones make sure to mark the front of the tissue block for better orientation and reproducibility of sections. Place the organ in the empty embedding mold in the appropriate orientation (see organ-specific details below) and fill with OCT until the organ is covered. Add an additional 1-2 mm of OCT on top of the organ for cryostat adjustments. 3. Organs are always frozen in the same orientation for reproducibility. a. trachea, the end that was close the thymus is facing downwards. b. thorax package, with thymus facing upwards. c. intestine, you can either freeze each subcompartment in one block if you have multiple animals or the four different parts in one block for each animal. We recommend to note in which corner you placed which tissue piece. d. tongue, the tip of the tongue facing upwards. This way you will reach the taste buds early during sectioning. e. nose, the tip of the nose is facing upwards. f. brain, the bulb is facing upwards and the hypothalamus is facing the front of the embedding mold. 5. Place the staining glass in a second larger container and surround it with dry ice. Fill the glass with isobutane around 0.5 to one cm high. Add ethanol to the dry ice. Make sure that the ethanol level is higher than the isobutane level.

Step 2.2: Preparation of cryosections
We recommend to section all organs in series of five. This means that section number two on slide number one is the sixth section in the series. This method was originally established from detailed brain mapping as this way the distance between one section to the next on the same slide is roughly the same as the distance represented from one brain atlas picture to the next. We define the section with the same position on subsequent slides as equal. This way you can do multiple stainings on equal sections. All organs are sectioned with a thickness of 10 μm, with the exception of the brain and pituitary which are sectioned with 14 μm. We highly recommend the use of disposable blades and a brush over the use of normal blades and a glass extender for sectioning.
For a general mapping, staining and imaging of representative sections from one series out of five is enough for each organ.
It is important to add the carrageenan first and then heat up. Let it stir between 37 °C and 40 °C overnight. Do not let the solution get warmer than 40 °C. On the following day, filter the carrageenan by gravity through filter paper and store at 4 °C. Volumes of up to 1 liter can be made as the buffer is stable at 4 °C.

Dissolve your antibodies in PBS-Carrageenan and fill the staining container with your antibody
solution. Our recommended containers have a capacity of 10 slides and need 38-40 ml of antibody solution to cover the sections. You can reuse these solutions many times. 3. Working solution of our antibodies: 1:10 0 0 for chicken anti-GFP (primary) and 1:500 for goat antichicken 488 (secondary). We recommended to titrate the primary to determine the best dilution. We recommend that secondary antibodies are diluted 1:500. 4. After the initial dehydration step do not let the slides dry out at all. We do all washing steps with gentle agitation.
Detailed staining protocol: a. Take the slides out of -80 °C and let them dry for 30 min at room temperature (RT). After this step, slides should not be allowed to dry out. b. Rehydrate the slides by washing three times for five minutes in PBS. c. Block the sections in blocking solution for 1 h at room temperature. For our blocking solution, we also use the container system enabling this buffer to be reused multiple times. When staining results worsen or the solution exhibits a color change, the blocking solution should be exchanged. d. Transfer the slides from the blocking container into the primary antibody container and incubate overnight at 4 °C. Different primary antibodies can be mixed in one container and incubated together. If subsequent incubations are instead used, wash then with 1x PBST three times for five minutes with agitation between the incubation steps. e. In general, always wash your slides at least three times for five minutes at RT with agitation between incubation steps. f. Incubate your slides in secondary antibody for two hours at RT. g. Wash three times for five minutes in PBST with agitation. h. Incubate the slides in Hoechst solution for five to ten minutes at RT when additional nuclear stain is desired. i. Coverslip all sections with mounting medium.
For imaging, we highly recommend a fluorescent microscope which is capable of producing tile images in order to image the whole, or the majority of the section in one picture to give an overview of each organ. We use the AxioScan slidescanner from Zeiss to automate the imaging process. After imaging we postprocess the pictures and pick out representative images for each organ, sex and age to upload into our database.

Enrichment of proteins from mouse tissue by antibodies and detection by mass spectrometry
For the enrichment of a protein, specific antibodies can be either covalently immobilized to, for example, tosyl-activated affinity magnetic beads (a), or monoclonal or polyclonal antibodies can be bound to protein A/G magnetic beads (b). The later interaction between antibody and protein A/G magnetic beads is not covalent. The covalent immobilization of antibodies to magnetic beads can be undertaken prior to the planed enrichment experiment and these are stable at 4 °C for several months. It is recommended that tissues or cells to be used for the protein enrichment experiment, are collected at an earlier date, snap frozen in liquid nitrogen then stored at -80 °C. Frozen or fresh protein lysates are prepared by disruption in the presence of a detergent-containing buffer (e.g RIPA buffer). After ultracentrifugation, the protein lysates are used for incubation with the immobilized antibodies [3,4] .
Step 1: Covalent coupling of monoclonal-or polyclonal antibodies to tosylated magnetic beads Material • Dynabeads M-280 tosylactivated (ThermoFisher, Invitrogen, No. 142.04) • magnetic rack (for 1.5 ml reaction tubes, ThermoFisher) • affinity-purified antibodies (polyclonal antibodies should be purified against their antigenic peptide or protein, monoclonal antibodies from mouse for example can be purified from cell culture medium with protein A/G-agarose. The antibody concentration should be 1 mg/ml in 100 mM HEPES pH 7.6 or in coupling buffer B (see below.) If the concentration is too low or the antibody is not in the appropriate buffer, the solution should be ultrafiltrated and concentrated (for example with Vivaspin MW 50 0 0 0 or 10 0 0 0 0, or Millipore) , collect the beads and discard the supernatant as previously described. 9. Wash twice with 1 ml buffer E, dissecting the supernatant after each as previously described.
11. Store antibody-coupled beads at 4 °C until use (antibody column is stable for at least 3-6 months).
Step 2: Binding of monoclonal-or polyclonal antibodies to protein A/G magnetic beads Material • Protein A and Protein G magnetic beads (e.g. Pierce Protein A/G Magnetic Beads, ThermoFisher) • magnetic rack (for 1.5 ml reaction tube, ThermoFisher) • RIPA buffer: 150 mM NaCl, 50 mM Tris HCl, pH 8.0, 5 mM EDTA, 1% Nonidet P40, 0.1% SDS, 0.5% Na-deoxycholate, pH 7.4 including protease inhibitors (Complete, Mini Protease Inhibitor Cocktail, Roche) 1. Pipette 50 μl of protein A/G beads into a 1.5 ml reaction tube, collect the beads with a magnetic rack, discard the supernatant. 2. Wash beads with 1 ml RIPA buffer, collect the beads as previously described and discard the supernatant. 3. Add 10 μg of monoclonal or polyclonal purified anti-TRPV6 antibodies. 4. Incubate antibody with magnetic beads for 1 h at room temperature. 5. Collect the beads with a magnetic rack, discard the supernatant. 6. Wash antibody beads with 1 ml RIPA buffer, collect the beads with a magnetic rack, discard the supernatant. 7. Proceed directly with tissue protein lysate incubation (see below) or store antibody-beads at 4 °C until use. Do not freeze the antibody-beads.
Step 3: Preparation of mouse tissue protein lysates Material • 10 0-50 0 mg fresh or frozen (-80 °C) mouse tissue Repeat homogenisation with glass teflon potter. 5. Ultracentrifuge tissue homogenate for 45 min at 10 0 0 0 0 xg at 4 °C. 6. Transfer supernatant into fresh falcon tube. 7. Incubate 250 μl of tosylatedantibody magnetic beads or 10 μg polyclonal/or monoclonal antibody bound to protein A/G magnetic beads with entire tissue protein lysate for 12-16 h at 4 °C. 8. Collect magnetic beads with a magnet rack, discard the supernatant. 9. Wash the antibody columns five times with 1 ml RIPA buffer (including protease inhibitors) using the magnetic rack. 10. After the last washing step, remove washing solution from the magnetic beads very carefully. 11. Elute antibody complex by adding 50 μl denaturing buffer and incubate at 60 °C for 20 min. 12. Store elute at -20 °C until gel electrophoresis can be performed.