The following workflow describes the manual processing of formaldehyde-fixed, paraffin-embedded tissue sections for small-scale analyses to meet basic research needs. Using low-capacity staining containers (~50-60 ml reagent volume), it is assumed that no more than 10 slides holding 10-20 sections (1-2 sections/slide), are handled simultaneously. If required or available, the protocol can be adjusted to fully or semi-automated staining systems. However, given the complex nature of teratoma samples and the time required for microscopy and image taking, it is reasonable to limit the number of analysed specimens to a maximum of around six per run. Cyclophilin A stainings and respective reagent controls (e.g. isotype control) are performed on serial sections of the same tissue mounted onto one slide to sustain identical experimental conditions. Tissues from hESC- and control (Matrigel)-injected mice are never run separately but always processed in parallel.
Deparaffinization and rehydration of tissue sections, Timing ~1 h 40 min
For this protocol, formaldehyde-fixed, paraffin-embedded tissues (Box 2) are sectioned at 2-5 μm, mounted onto positively charged slides and dried overnight at room temperature in an upright position to allow sections to firmly adhere. Slides stored in a slide box at ambient temperature are then typically processed within 3-5 days. The effect of prolonged slide storage on staining quality is generally quite variable and largely depends on the individual protein, and thus has to be established empirically. In our experience, singleplex stainings of cyclophilin A can be performed on slides stored at room temperature for up to six months, with only minor to moderate loss in overall signal intensity. However, for any systematic comparisons, either qualitative or quantitative, and especially if co-staining for other proteins is performed, it is best to use freshly sectioned tissue stored for less than one week.
1 Pre-heat a Coplin staining jar (glass-based variant) in an incubator at 60°C for ~30 min. Do not screw on the lid.
2 Load jar with slides positioned upright. Place slides either individually or arrange them back-to-back, and incubate for another 30 min at 60°C to allow the paraffin wax to melt.
CRITICAL STEP This is a preparative step to soften the paraffin wax and to prime tissue sections for deparaffinization in xylene. It supports the efficient removal of paraffin which is essential to achieve a good signal-to-noise ratio in subsequent immunostaining. Although most protocols do not include a "slide baking step" prior to solvent-based dewaxing, this approach has proven extremely useful in the context of the procedure and thus should not be omitted.
All subsequent washes are performed at room temperature under constant, mild agitation on an orbital shaker. Make sure that slides are fully immersed to compensate for varying reagent levels during rotation. Do not submerge the slide grip (~30-40 ml reagent volume/staining jar).
3 After slide baking, immediately proceed to deparaffinization by incubating sections in two washes of xylene, 10 min each.
CAUTION Xylene is volatile and toxic, and should be handled carefully. Perform steps in a chemical fume hood or screw on the jar cap to reduce solvent evaporation and to avoid spill during incubation. For re-use, store tightly capped in a fume hood.
CRITICAL STEP Xylene solutions are typically re-used but will exhaust over time and gradually loose the ability to dissolve the paraffin wax completely. Make sure to replace solutions on a regular basis, depending on the sample throughput (e.g. once every 60-120 sections). Streaking is highly indicative of exhausted solutions which should be replaced with fresh xylene immediately. Insufficient removal of paraffin will significantly impact overall staining quality, mostly by causing uneven, spotty background staining and by generally decreasing the efficiency of antigen detection.
4 Using forceps, transfer slides into a staining jar with 2-propanol and incubate for 5 min. Make sure to carefully wipe off the xylene to minimize carryover.
CRITICAL STEP Propanol and anhydrous ethanol are considered to perform more or less equally well in tissue de- and rehydration, with propanol being cheaper and readily accessible but requiring prolonged incubation. This protocol employs propanol as an intermediate step to roughly clear most of the xylene before proceeding to 100% ethanol for ultimate removal. Additionally, unlike ethanol, propanol is somewhat miscible with molten paraffin which might favour wax removal. Overall, this step reduces the total amount of ethanol required, and has proven beneficial to the overall outcome of the procedure. Do not reduce incubation time.
5 For rehydration, transfer sections through a series of graded ethanol solutions, that is once through 100%, 70% and 50% ethanol, 2 min each.
CRITICAL STEP Xylene does not mix with water. For its efficient removal, always use ethanol with low water content (≤1%) (absolute grade ethanol). Flow marks indicate the use of inappropriate ethanol formulations. Propanol and ethanol solutions can be re-used but are exhanged on a daily basis in this protocol. If re-used, store in staining jars with the cap screwed on to minimize evaporation.
6 After rehydration, rinse sections three times in dH2O, 5 min each. Keep slides in dH2O until ready to perform antigen retrieval.
CRITICAL STEP Once the deparaffinization and rehydration steps have been completed, it is essential that sections do not dry out at any time during the subsequent procedure. Depending on the extent of drying, this will either prevent any specific staining with no or only very weak signals being detected, or promote non-specific antibody binding, causing extensive background staining.
Heat-induced epitope retrieval (HIER), Timing ~60 min
For this protocol, antigen unmasking for subsequent immunostaining has been established using a domestic, 800-watt microwave oven of standard quality. It is operated manually in a discontinous mode by following a defined sequence of on/off cycles at two different power levels. If using a scientific microwave with more even heating and especially accurate temperature and timing control, prior testing might be required to convert the here specified parameters into instrument settings that perform equally well.
7 While performing step 6 of the procedure, prepare a sufficient amount of 1X antigen retrieval buffer and pre-warm in an incubator at 60°C until ready to proceed (~10-15 min).
CRITICAL STEP For best results, prepare 1X solutions freshly right before use or on a daily basis. Make sure to include some extra buffer to compensate for evaporation or boilover during microwaving, altogether ~50 ml retrieval buffer per staining jar.
8 Transfer ~40 ml pre-heated 1X antigen retrieval buffer into a microwaveable staining jar (e.g. plastic Coplin staining jar). Save the rest.
CAUTION For reasons of safety, always use vessels or racks made of microwave-proof plastic (e.g. polypropylene or high-density polyethylene). Standard histology glass is heated more unevenly and will mostly crack when microwaved. It should be noted however, that during the set-up of the procedure, the heating protocol outlined below could successfully be completed using the glass Coplin jars specified in the Materials section. For this, only use intact jars with no visible signs of glass damage or fatigue, and allow the heated jar to rest in the oven for ~1-2 min before taking it out to proceed. This minimizes the likelihood of cracking by avoiding a sudden drop in temperature. Do not use these jars in combination with any other heating protocols than the one specified in this procedure.
9 Remove slides from water (step 6) and immediately place in retrieval buffer. Make sure that slides are covered with buffer by at least one centimeter, that is fully immersed including half of the slide grip, to allow for evaporation during boiling. Do not screw on the jar cap.
10 Center the jar inside the microwave and adjust the power level to "low". For the device specified in this protocol, this level equals 10% of the total power, that is 80 W, and very roughly corresponds to a temperature of <100°C (~65-90°C).
11 Turn on the oven and wait until the solution comes to an initial boil, typically 3-6 min. Keep boiling for 3-5 sec and immediately turn off. Pause for 10 sec, with the door open.
CRITICAL STEP Microwaves in general, but especially those for domestic use, are known to produce uneven standing wave patterns, leading to the formation of so called hot and cold spots and thus uneven heating. This can cause unbalanced epitope retrieval and hence uneven staining due to a variance in antigen accessibility. For uniform epitope retrieval, always microwave with the rotation dish on to balance hot and cold spots during heating.
12 For non-continous heating at 25% power (200 W) and 90-110°C, switch to the "defrost" level.
13 Turn on and wait until boiling is initiated, typically <30 sec. Keep boiling for 1-2 sec and immediately turn off. Pause for 10 sec, with the door open.
CRITICAL STEP This procedure uses non-sealed vessels. While heating, be sure to monitor the buffer level in the jar and to watch out for excessive evaporation or boilover. Fill up with retrieval buffer if necessary. Do not allow the slides to dry out at any time during microwaving.
14 Repeat on/off cycles (step 13) for ~2-4 times.
CRITICAL STEP For best results, follow the antigen retrieval protocol as outlined above. Importantly, these conditions have been proven to work well in both, cyclophilin A singleplex and double staining approaches. Do not reduce the number of boiling steps as this might leave the antigen(s) largely un-retrieved and thus poorly accessible for detection. Excessive boiling on the other hand, either due to multiple boiling steps or an increase in power/temperature, might result in over-retrieved epitopes, leading to high background staining. But more often, rigorous boiling significantly increases the chance of sections dissociating from the slide, and thus should be largely avoided.
15 After the last boiling step has been completed, remove the staining jar from the microwave and allow the slides to cool to room temperature on benchtop for ~30 min.
CAUTION Be careful when handling hot liquids.
CRITICAL STEP Do not omit this step to directly proceed to immunostaining. Cooling slides to room temperature does not only permit to handle them properly, but allows the antigens to recover from heating and importantly, reduces the likelihood of sections coming off during the subsequent staining procedure.
16 To eliminate the retrieval buffer and to prepare sections for immunostaining, transfer slides into a staining jar with 1X DPBS and rinse three times, 5 min each.
Immunostaining, Timing ~20 h
All subsequent steps are performed at room temperature unless otherwise noted. The blocking and antibody incubation steps are routinely carried out in a light-proof moisture chamber to prevent tissues from drying, and to protect fluorophores from light. During the set-up of the procedure, two different blocking agents have been tested for their ability to reduce non-specific binding, i.e. bovine serum albumin (BSA) (option A) and normal (unchallenged) serum from the host species that the secondary antibody was raised in (normal goat serum) (option B). In our hands, either condition consistently yields a good signal-to-noise ratio in both, cyclophilin A singleplex as well as double staining approaches. In summary, both options give very comparable results, making them virtually interchangeable in this procedure.
17 While performing steps 15 and 16 of the procedure, prepare an appropriate amount of blocking solution (option A or B), enough to perform the subsequent blocking and primary antibody incubation steps (~200-400 μl/tissue section). Tissue sections processed in this protocol have an average size of 1.0-1.5 cm2 and are typically covered with 100-200 μl reagent volume. 10 ml blocking solution is therefore sufficient to complete the maximum amount of 20 sections per run, including some extra solution to compensate for pipetting errors or other mistakes. For optimum results, prepare freshly right before use or on a daily basis.
CRITICAL STEP Detergent-mediated permeabilization can greatly improve antibody access to antigens inside the cell, including nuclear proteins. Moreover, the addition of a harsh detergent such as Triton X-100 might also help to decrease the general background by clearing the tissue from potential non-specific binding sites. The blocking solution employed in this protocol contains 0.3% (v/v) Triton-X100, and has been optimized for the detection of cyclophilin A and other intracellular proteins, e.g. α-smooth muscle actin (SMA) or phosphorylated S6, and has been demonstrated to perform equally well for the adhesion molecule CD31. It should be noted however, that these conditions might be too harsh if co-staining of other cell surface epitopes is required, as those might get disrupted or partially dissolved, and consequently loose their antigenicity. For this, Triton X-100 could be exchanged for a somewhat milder detergent such as Tween 20 or saponin, but this has to be tested empirically.
18 Humidify the moisture chamber by adding dH2O to the bottom of each slide compartement (~6-7 ml). Refer to the manufacturer’s instructions for detailed information on how to humidify your incubation box.
For subsequent steps, process slides one after another to prevent them from drying-out.
19 Remove slides from 1X DPBS (step 16) and quickly drain on a paper towel to remove most of the excess liquid. Do not allow slides to dry completely.
20 Using a liquid-repellent slide marker, draw a narrow circle around the tissue section and wait for ~5-10 sec.
CAUTION The pen contains hazardous ingredients including bromopropane and ligroin (a fraction of petroleum). Avoid contact with skin and eyes and wear gloves.
CRITICAL STEP Liquid-repellent slide markers can be used to draw hydrophobic barriers on glass slides that confine the flow of a reagent to a defined area, providing a water-resistant containment to which various reagents can be applied. They are extremely useful in this procedure and allow to independently process two sections on the same slide (e.g. specific staining and respective reagent control). Moreover, they help to save precious reagents and to locate the tissue after mounting. It should be noted however, that it is essential that slides are not allowed to dry out during any time of the staining procedure. For the liquid blocker to be applied, slides need not to be dry, and the markings can be well drawn on wet slides. The hydrophobic properties of the barrier will force the residual 1X DPBS on the slide towards the center of the encircled area, preventing the tissue from drying out. Markings are insoluble in aqueous buffers, detergents, alcohol and acetone but can be removed with xylene, if necessary. Shake pen well before use.
21 Shake off any residual 1X DPBS from the slides and transfer them to the moisture chamber. If excessive amounts of 1X DPBS are left, especially on top of the sections, carefully pipette them off. Do not allow the tip to touch the tissue.
22 Immediately apply ~100-200 μl blocking solution (option A or B) to the sections. Add dropwise. Make sure to not only cover the tissue itself but the entire area encircled by the hydrophobic barrier. If applied correctly, the liquid will form a dome-shaped small heap within the markings.
CRITICAL STEP For teratoma tissue engrafted into mouse skeletal muscle, blocking option A and B perform equally well in our hands, and are thus mutually exchangeable. This might also be due to the fact that monoclonal primaries are used which in general show reduced cross-reactivity towards unrelated epitopes. If other tissues are injected, or if double stainings with polyclonals are planned, option B might give better results but this has to be tested empirically.
23 Incubate for 1 h at room temperature.
24 While blocking, prepare primary antibody and isotype control.
A. First option: singleplex staining (cyclophilin A only)
i. Dilute Cyclophilin A (D2Y4M) rabbit monoclonal antibody in blocking solution (option A or B) to a final concentration of 1.3 μg/ml. Mix gently by inverting and spin down briefly. Keep at 4°C until ready to proceed.
CRITICAL STEP For optimum results, do not switch the type of blocking solution between blocking and antibody incubation steps, i.e. option A to B or B to A. The given working concentration is based upon antibody titration to obtain a high signal-to-noise ratio in cyclophilin A single and double stainings. It lies within the range of commonly applied concentrations (~0.5-10 μg/ml, occasionally up to 25 μg/ml) but at the lower end, with relatively small amounts of antibody being sufficient to achieve a good staining. For lot #1 of the antibody, which has been used throughout, this working concentration corresponds to a dilution of 1:100, i.e. 1-2 μl cyclophilin A antibody per 100-200 μl blocking solution per tissue section. At Cell Signaling Technology, antibody concentrations are lot-specific and may vary between two different lots of the same antibody. Hence, if using a different lot, do not dilute according to the ratio notation but always refer to the antibody concentration of the respective lot to calculate the amount of antibody actually required. Lot-specific antibody concentrations are available upon request ([email protected]). Be aware that knowledge of the actual working concentration is a prerequisite for the correct application of isotype controls. For best results, stick to the conditions specified above. No further optimization of antibody concentration is required.
ii. Pre-dilute rabbit (DA1E) monoclonal IgG (isotype control) 1:20 in 1X DPBS to a concentration of 125 μg/ml, and further dilute in blocking solution (option A or B) to a final concentration of 1.3 μg/ml. This corresponds to 1.04-2.08 μl pre-diluted rabbit IgG per 100-200 μl blocking solution per tissue section. Mix gently by inverting and spin down briefly. Keep at 4°C until ready to proceed. Pre-diluted IgG can be aliquoted into working volumes and stored at -20°C for further use. Thaw only once, with the remainder kept at 4°C for ~1-2 weeks.
CRITICAL STEP Always make sure that the primary antibody and the respective isotype control are concentration- (not dilution-) matched. Incorrect dilution of the isotype control can cause either false-positive or false-negative results. Pre-dilution of the control IgG is recommended to avoid inaccuracy through pipetting errors. For lot-independent concentration of the rabbit IgG, see the datasheet or the supplier’s homepage (https://www.cellsignal.com/)
B. Second option: double staining (cyclophilin A and SMA), simultaneous protocol
i. Prepare cyclophilin A antibody as described above (step 24, A, i) and add SMA (1A4) mouse monoclonal antibody to a final concentration of 10 μg/ml. This corresponds to 1-2 μl SMA antibody per 100-200 μl cyclophilin A antibody-containing blocking solution per tissue section. Mix gently by inverting and spin down briefly. Keep at 4°C until ready to proceed.
ii. Prepare rabbit isotype control as described above (step 24, A, ii) and add mouse (MOPC-173) monoclonal IgG2a (isotype control) to a final concentration of 10 μg/ml. This corresponds to 2-4 μl mouse IgG2a per 100-200 μl rabbit IgG-containing blocking solution per tissue section. Mix gently by inverting and spin down briefly. Keep at 4°C until ready to proceed.
25 After blocking is completed, remove slides from the moisture chamber and quickly tilt, long edge down, to drain off the blocking buffer. Using a small piece of paper towel, carefully blot off the liquid next to the hydrophobic barrier. Do not allow slides to dry completely.
26 Put slides back in the moisture chamber and immediately apply 100-200 μl diluted primary antibody or isotype control to the sections. Make sure to use sufficient volumes. To protect against spill-out during long-term incubation, a second hydrophobic barrier can be drawn right next to the first one. Do so before antibody solutions are applied.
27 Place the moisture chamber in a fridge or cold room, and incubate overnight (~16 h) at 4°C.
CRITICAL STEP The procedure has been optimized for a low cyclophilin A antibody titer and overnight incubation to achieve a slow but targeted binding and to reach a saturation point as possible where all antigen is bound. Incubation periods up to 24 h consistently yield good staining results. Do not reduce incubation time.
28 On the next day, prepare a sufficient amount of blocking solution (option A or B) (~100-200 μl/tissue section). Stick to the option used the day before.
29 Remove slides from the moisture chamber and drain off the antibody solution on a paper towel. If slides are mounted with two sections that are incubated with different antibodies and/or control reagents, take care to tilt them against their long edge to prevent solutions from blending into each other.
30 Transfer slides into a staining jar with 1X DPBS and wash three times, 5 min each.
31 While washing, prepare fluorophore-conjugated secondary antibody.
A. First option: singleplex staining (cyclophilin A only)
i. Dilute Alexa Fluor 488-conjugated anti-rabbit IgG 1:500 in blocking solution (option A or B) to a final concentration of 4 μg/ml. This corresponds to 0.2-0.4 μl anti-rabbit IgG per 100-200 μl blocking solution per tissue section. Mix gently by inverting and spin down briefly. Protect from light until ready to proceed.
B. Second option: double staining (cyclophilin A and SMA), simultaneous protocol
i. Prepare Alexa Fluor 488-conjugated anti-rabbit IgG as described above (step 31, A, i) and add Alexa Fluor 594-conjugated anti-mouse IgG at a dilution of 1:500 to yield a final concentration of 4 μg/ml. This corresponds to 0.2-0.4 μl anti-mouse IgG per 100-200 μl anti-rabbit IgG-containing blocking solution per tissue section. Mix gently by inverting and spin down briefly. Protect from light until ready to proceed.
CRITICAL STEP The staining of mouse tissue using a mouse primary antibody is often associated with increased levels of background staining, mainly caused by the secondary antibody binding to endogenous mouse IgG or Fc receptors on tissue cells (e.g. macrophages). The latter can be largely eliminated by using F(ab')2 fragment antibodies as outlined in this procedure. See also the Materials section. The remainder of background staining is usually fixed via a separate blocking step, mostly by using commercially available kits. At this point however, it has to be noted that the mouse-on-mouse detection using this protocol (e.g. SMA or phosphorylated S6 on mouse cells/ mouse tissue)7 did not result in a decreased signal-to-noise ratio. However, if other tissues are injected, an additional blocking step might be required.
32 After the last washing step is completed, remove slides from 1X DPBS and shake off excess liquid. Place slides in the moisture chamber and immediately add 100-200 μl diluted fluorophore-conjugated secondary antibody onto the sections.
33 Incubate for 1 h at room temperature.
34 Remove slides from the moisture chamber and drain off the antibody solution on a paper towel. Transfer slides into a staining jar with 1X DPBS and wash three times, 5 min each.
CRITICAL STEP Thorough washing of sections between and after antibody incubation steps is crucial to minimize background staining and to contrast the specific signal. For optimal results, do neither reduce the number nor the duration of washing steps.
35 While washing, prepare an appropriate amount (~30-40 ml) of DAPI working solution and transfer into a staining jar.
36 Remove slides from 1X DPBS and quickly drain on a paper towel. Counterstain sections by immersing slides for 3-5 min. Alternatively, 100-200 μl DAPI solution can be directly applied to the sections. Do not incubate for more than 5-8 min. The DAPI solution can be re-used for several times when stored at 4°C. However, for consistent results prepare solutions freshly right before use.
37 Carefully drain off the DAPI solution and rinse slides once in 1X DPBS for 5 min. Keep slides in 1X DPBS until ready to perform coverslipping.
38 Apply ~4-8 20 μl-drops of antifade mounting medium directly to a coverslip.
39 Remove slides from 1X DPBS and quickly drain on a paper towel. Eliminate residual liquid by vigorously shaking it off. Lay slides, section side down, over the coverslip. Absorb excess mounting medium with a paper towel.
CRITICAL STEP Store mounted slides flat or on the edge at 4°C protected from light, e.g. in a slide folder or slide box, until further processed. Sections are ready for viewing and image taking right after coverslipping but for best results, store overnight (8-12 h) at 4°C. Cyclophilin A-stained samples are typically assessed within 2-5 days but might be stored longer (2-3 weeks). The aqueous mounting medium is non-hardening, i.e. it remains liquid on the slide and does not solidify. Mounted slides are usually stored without sealing. However, for long-term storage, coverslips can be sealed with nail polish or an appropriate sealant.
40 Subject slides to microscopic evaluation.
CRITICAL STEP For optimum results, run a pilot experiment with a limited number of samples to establish an appropriate imaging protocol which can be used throughout experimentation. This protocol typically includes parameters such as excitation intensity, detector gain and exposure time, and many more. If available, a transmitted light imaging mode (e.g. phase contrast) should be incorporated into these settings. The phase contrast and confocal epifluorescence images can be acquired simultaneously, and when merged, give valuable information about the precise location of the labeled cells within the tissue. This feature is especially helpful for studies of the teratoma/tumor microenvironment since it allows to track host cell infiltration, and at the same time to evaluate its impact on tissue properties and condition. Always use the same imaging and post-acquisition protocols to analyse your samples of interest and the respective positive and/or negative controls.