The answers to many questions in biology lie in the ability to examine the relevant biological structures accurately at high resolution. Electron microscopy (EM) offers the unparalleled power to study cellular morphology and structure at nanoscale resolution (Leapman, 2004). Cryofixation by high-pressure freezing (hereafter referred to as cryofixation) is the optimal fixation method for samples of thicknesses up to approximately 500 µm (Dahl and Staehelin, 1989; McDonald, 1999; Moor, 1987; Shimoni et al., 1998). By rapidly freezing the samples in liquid nitrogen (−196 °C) under high pressure (~2100 bar), cryofixation immobilizes cellular structures within milliseconds and preserves them in their near-native state. In contrast, cross-linking-based chemical fixation takes place at higher temperatures (≥4 °C) and depends on the infiltration of aldehyde fixatives, a process which takes seconds to minutes to complete. During chemical fixation, cellular structures may deteriorate or undergo rearrangement (Korogod et al., 2015; Steinbrecht and Müller, 1987; Szczesny et al., 1996) and enzymatic reactions can proceed (Kellenberger et al., 1992; Sabatini et al., 1963), potentially resulting in significant morphological artefacts.

Cryofixation is especially critical, and often necessary, for properly fixing tissues with cell walls or cuticles that are impermeable to chemical fixatives, such as samples from yeast, plant, C. elegans, and Drosophila (Ding et al., 1993; Doroquez et al., 2014; Kaeser et al., 1989; Kiss et al., 1990; McDonald, 2007; Müller-Reichert et al., 2003; Shanbhag et al., 1999; Shanbhag et al., 2000; Winey et al., 1995). As cryofixation instantaneously halts all cellular processes, it also provides the temporal control needed to capture fleeting biological events in a dynamic process (Hess et al., 2000; Watanabe et al., 2013a, 2013b, 2014b).

Despite the clear benefits of cryofixation, it is incompatible with diaminobenzidine (DAB) labeling reactions by genetic EM tags. For example, APEX2 (enhanced ascorbate peroxidase) is an engineered peroxidase that catalyzes DAB reaction to render target structures electron dense (Lam et al., 2015; Martell et al., 2012). Despite the successful applications of APEX2 to three-dimensional (3D) EM (Joesch et al., 2016), there has been no demonstration that APEX2 or other genetic EM tags can be activated following cryofixation. Conventionally, cryofixation is followed by freeze-substitution (Steinbrecht and Müller, 1987), during which water in the sample is replaced by organic solvents. However, the resulting dehydrated environment is incompatible with the aqueous enzymatic reactions required for DAB labeling by genetic EM tags.

EM structures can also be genetically labeled with fluorescent markers through correlated light and electron microscopy (CLEM). Yet, performing CLEM with cryofixed samples also presents challenges. Fluorescence microscopy commonly takes place either before cryofixation (Brown et al., 2009; Kolotuev et al., 2009; McDonald, 2009) or at a later stage after the sample is embedded (Kukulski et al., 2011; Nixon et al., 2009; Schwarz and Humbel, 2009). However, if the specimen is dissected from live animals, the time taken to acquire fluorescence images delays cryofixation and could cause ultrastructural deterioration. In order for fluorescence microscopy to take place after embedding, special acrylic resins need to be used (Kukulski et al., 2011; Nixon et al., 2009; Schwarz and Humbel, 2009) and only a low concentration of osmium tetroxide stain can be tolerated (de Boer et al., 2015; Watanabe et al., 2011). Although one can in principle perform fluorescence microscopy in cryofixed samples after rehydration, fluorescence images have only been acquired in sucrose-infiltrated cryosections (300–500 nm) (Ripper et al., 2008; Stierhof and El Kasmi, 2010). Moreover, no protocol has been developed to prepare large cryofixed tissues expressing genetic CLEM markers for high-contrast EM imaging. These constraints limit the applicability of CLEM for cryofixed samples.

Another disadvantage of cryofixation is that en bloc staining during freeze-substitution is often inadequate. As a result, post-staining of ultramicrotomy sections is frequently needed for cryofixed samples (Shanbhag et al., 1999; Shanbhag et al., 2000; Takemura et al., 2013). However, post-staining could be labor-intensive and time-consuming, especially for volume EM (Ryan et al., 2016; Zheng et al., 2017). Critically, on-section staining is impossible for samples imaged with block-face volume EM techniques (Briggman and Bock, 2012), such as serial block-face scanning electron microscopy (SBEM) (Denk and Horstmann, 2004). A large amount of heavy metal staining is necessary for SBEM to generate sufficient back-scatter electron signal and to prevent specimen charging (Deerinck et al., 2010; Kelley et al., 1973; Tapia et al., 2012). Therefore, it remains impossible to image cryofixed samples with SBEM or other techniques that require high-contrast staining.

To overcome these limitations of cryofixation, here we present a robust approach, named the CryoChem Method (CCM), which combines key advantages of cryofixation and chemical fixation. This technique enables labeling of target structures by genetically encoded EM tags or fluorescent markers in cryofixed samples, and permits high-contrast en bloc heavy metal staining sufficient for SBEM. Specifically, we rehydrate cryofixed samples after freeze-substitution to make the specimen suitable for subsequent aqueous reactions and fluorescence imaging. We successfully apply CCM to multiple biologically significant systems with distinct ultrastructural morphology, including cultured mammalian cells, Drosophila olfactory organ (antenna) and mouse brain. By overcoming critical technical barriers, our method exploits the potential of cryofixation, making it compatible with genetically encoded EM tags and any EM techniques that require substantial heavy metal staining. Furthermore, the versatility of CCM allows us to achieve 3D CLEM in a well-preserved mouse brain by permitting SBEM after fluorescent imaging of a frozen-rehydrated specimen.

Given that a key limitation of cryofixation arises from the dehydrated state of the samples after freeze-substitution (Table 1), it is imperative that our approach delivers a cryofixed specimen that is fully hydrated and can then be processed at higher temperatures (4 °C or room temperature) for enzymatic reactions and/or high-contrast en bloc heavy metal staining. It has been demonstrated that cryofixed samples can be rehydrated for immunogold labeling or fluorescence imaging following cryosectioning (Dhonukshe et al., 2007; Ripper et al., 2008; Stierhof and El Kasmi, 2010; van Donselaar et al., 2007), but these approaches only yield modest EM contrast. In addition, the methods are incompatible with volume EM techniques and have yet to be successfully combined with genetic labeling using APEX2.

The CryoChem method

To achieve the ultrastructural preservation of cryofixation and the versatility of chemical fixation, we developed a hybrid protocol which we refer to hereafter as the CryoChem Method (CCM) (Table 1). Importantly, we devised a freeze-substitution cocktail (see below) that allows preservation of APEX2 enzymatic activity and signals from fluorescent proteins. CCM begins with high-pressure freezing of a sample, followed by freeze-substitution in an acetone solution with glutaraldehyde (0.2%), uranyl acetate (0.1%), methanol (2%) and water (1%), to further stabilize the cryo-preserved structures at low temperatures. After freeze-substitution, the sample is rehydrated gradually on ice with a series of acetone solutions containing an increasing concentration of water or 0.1 M HEPES. Once completely rehydrated, the cryofixed sample is amenable for imaging with fluorescence microscopy, DAB labeling reactions using genetically encoded tags, and the high-contrast en bloc staining (e.g. osmium-thiocarbohydrazide-osmium and uranyl acetate) normally reserved only for chemically fixed samples. Afterwards, the samples is dehydrated through a series of ethanol solutions and acetone, then infiltrated with epoxy resin and cured using standard EM procedures. To minimize volume artefact, epoxy resin is chosen because it causes minimal tissue shrinkage during embedding (<2%) compared to other embedding media (Kushida, 1962). The resin-embedded sample can be sectioned or imaged directly with any desired EM technique (Figure 1, see Materials and methods for details).

Figure 1

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CryoChem method offers high-quality ultrastructural preservation and sufficient en bloc staining for SBEM

To determine whether CCM provides high-quality ultrastructural preservation, we first tested the method in a mammalian cell line. Using transmission electron microscopy (TEM), well-preserved mitochondria and nuclear membrane were observed in the CCM-processed cells (Figure 2—figure supplement 1). Given that cryofixation is often necessary for properly fixing tissues surrounded by a barrier to chemical fixatives (Steinbrecht, 1980; Steinbrecht and Müller, 1987), we next tested CCM in a Drosophila olfactory organ, the antenna, which is encased in a waxy cuticle (Figure 2A and B). A hallmark of optimally preserved antennal tissues prepared by cryofixation is the smooth appearance of membrane structures (Shanbhag et al., 1999; Shanbhag et al., 2000; Steinbrecht, 1980; Steinbrecht and Müller, 1987). In the insect antenna, auxiliary cells extend microlamellae to surround the olfactory receptor neurons (ORNs), forming the most membrane-rich regions in the antenna. We therefore focused on this structure to evaluate the quality of morphological preservation afforded by our method. In the CCM-processed antennal tissues, we found that the delicate structure of the microlamellae was well-preserved (Figure 2B and Figure 2—video 1), unlike the chemically fixed counterparts in which microlamellae were disorganized and distorted (Figure 2A) (Steinbrecht, 1980). Furthermore, there were numerous signs of extraction of cellular materials in the chemically fixed antenna (Figure 2A1, arrows), but not in the CCM-processed specimen (Figure 2B). Importantly, the overall ultrastructural preservation achieved through CCM resembles that obtained by standard cryofixation and freeze-substitution protocols (Shanbhag et al., 1999; Shanbhag et al., 2000). This observation also suggests that the rehydration step in CCM leads to little, if any, swelling in the antenna tissue.

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In contrast to fly antennae, which can be dissected expeditiously and frozen in the live state, certain tissues (e.g., mouse brain) are difficult to cryofix from life without tissue damage caused by anoxia or mechanical stress associated with dissection. In these cases, cryofixation can be performed after aldehyde perfusion and still produce quality morphological preservation (Sosinsky et al., 2008). To test whether CCM can improve morphological preservation of aldehyde-perfused samples, we cryofixed vibratome sections (100 µm) from an aldehyde-perfused mouse brain and processed the sample with CCM. Compared to specimens processed by a standard EM preparation method that involved dehydration on ice (Figure 2C), the CCM-processed samples, which were initially dehydrated through freeze-substitution, showed smoother membranes and an increase in cytoplasmic density (Figure 2D). This result indicates an improvement in morphological preservation and agrees with our previous observation that cellular morphology can be markedly improved even when cryofixation is performed after aldehyde perfusion (Sosinsky et al., 2008).

Of note, we adopted a high-contrast en bloc staining protocol (Deerinck et al., 2010; Tapia et al., 2012; West et al., 2010; Williams et al., 2011) when processing the Drosophila antennae and mouse brain. An adequate level of heavy metals was incorporated into these cryofixed samples to allow for successful imaging by SBEM (Figures 2B and 3), even without nitrogen gas injection to dissipate any charge build-up that often occurs on samples of low conductivity (Deerinck et al., 2018) (Figure 2—video 1 and Figure 3D). This en bloc staining protocol is normally reserved only for chemically fixed tissues, but is now made compatible with cryofixed samples by CCM.

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Fluorescence is well-preserved in CryoChem-processed samples

To determine whether CCM is compatible with fluorescence microscopy, we first evaluated the degree to which fluorescence level is affected after CCM processing. Using confocal microscopy, we quantified GFP fluorescence in the soma of unfixed Drosophila ORNs and that from the CCM-processed samples after rehydration (Figure 4). Remarkably, GFP fluorescence intensities of the fresh and the CCM-processed ORNs were essentially indistinguishable with respect to their distributions (Figure 4A) and average levels (Figure 4B). This result indicates that CCM processing has little effect on GFP fluorescence in fly ORNs, likely due to the use of mild fixatives during freeze-substitution in our protocol. Similarly, we observed strong GFP signals in the mouse brain after the cryofixed sample was rehydrated (Figure 4—figure supplement 1A).

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Figure 4—source data 1

GPF fluorescence intensities in unfixed and CryoChem-processed Drosophila ORN soma.

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Next, we asked whether this observation also applies to another type of fluorescence protein. To this end, we examined tdTomato fluorescence in the mouse brain (Figure 4—figure supplement 1B). We note that tdTomato is not a variant of GFP and is instead derived from Discosoma sp. fluorescence protein ‘DsRed’ (Shaner et al., 2004). Confocal images of the CCM-processed mouse brain showed that the tdTomato fluorescence was also well-preserved (Figure 4—figure supplement 1B) and we were able to detect the co-expression of GFP and tdTomato in a subpopulation of neurons (Figure 4—figure supplement 1C). Together, our results indicate that CCM-processed sample can serve as a robust substrate for fluorescence imaging. As such, CCM allows fluorescence imaging to be combined with DAB labeling and high-contrast en bloc staining in the same cryofixed sample, a critical advance to cryofixation-rehydration methods (Dhonukshe et al., 2007; Ripper et al., 2008; Stierhof and El Kasmi, 2010; van Donselaar et al., 2007).

3D correlative light and electron microscopy (CLEM) in CCM-processed samples expressing fluorescent markers

We took advantage of the fact that fluorescence microscopy can take place in a cryofixed sample before resin embedding to develop a protocol for 3D CLEM in CCM-processed specimens (Figure 5A, see Materials and methods for details), so that the correlation can be achieved in optimally preserved tissues. The protocol first uses the core CCM steps to deliver a frozen-rehydrated sample. Subsequently, DRAQ5 DNA stain is introduced to the sample to label the nuclei, which can then serve as fiducial markers for CLEM. Next, the region containing target cells expressing fluorescent markers is imaged with confocal microscopy, during which signals from DRAQ5 and fluorescent markers are both acquired. After confocal microscopy, the sample is en bloc stained with multiple layers of heavy metals (Deerinck et al., 2010; Tapia et al., 2012; West et al., 2010; Williams et al., 2011), then dehydrated and embedded as in a typical CCM protocol. Subsequently, the embedded sample is imaged with X-ray microscopy. The resulting micro-computed tomography volume can be registered to the confocal volume using the nuclei as fiducial markers, so that the region of interest (ROI) for SBEM can be identified. After SBEM imaging, the EM volume can be registered to the confocal volume in a similar fashion for 3D CLEM.

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As a proof of principle, we performed 3D CLEM in an aldehyde-perfused, CCM-processed mouse brain expressing tdTomato in a subset of neurons. To this end, we first determined if DRAQ5 staining can be performed in a frozen-rehydrated specimen. Using confocal microscopy, we were able to observe DRAQ5 labeling of the nuclei in a cryofixed brain slice after rehydration (Figure 5B). We used the labeled nuclei as fiducial markers to register the X-ray volume with the confocal data (Figure 5B) and thereby target a ROI with tdTomato-expressing neurons for SBEM imaging.

Similarly, we were able to register the confocal volume to the SBEM volume (Figure 5C). Of note, the CLEM accuracy was ensured by using a subset of DRAQ5-labeled heterochromatin structures and their corresponding counterparts in EM as finer fiducial points (Figure 5C).

The fluorescent markers made it possible to identify the target cell bodies (Figure 5D and Figure 5—video 1) in the SBEM volume. The high accuracy of correlation achieved by our 3D CLEM protocol is demonstrated by the successful alignments of multiple ultrastructures: the fine neuronal processes (Figure 5E) and a subcellular heterochromatin structure that was not used as a fiducial marker (Figure 5—figure supplement 1). Lastly, we note that with CCM, fluorescence microscopy in cryofixed specimens takes place before en bloc EM staining. Therefore, our protocol does not require special resins for embedding and permits high-contrast staining with high concentrations of osmium tetroxide.

We described here a hybrid method, named CryoChem, which combines key advantages of cryofixation and chemical fixation to substantially broaden the applicability of the optimal fixation technique. With CCM, it is now possible to label target structures with DAB by a genetically encoded EM tag and deposit high-contrast en bloc staining in cryofixed tissues. In addition, with CCM, one can image cells expressing fluorescent markers before resin embedding and perform 3D CLEM in cryofixed specimens. Our method thereby provides an alternative to conventional cryofixation and chemical fixation methods.

The modular nature of CCM (Figure 1) makes it highly versatile as researchers can modify the modules to best suit their needs. For instance, to prevent over-staining, one can replace the high-contrast en bloc staining step (osmium-thiocarbohydrazide-osmium and uranyl acetate) (Deerinck et al., 2010; Tapia et al., 2012; West et al., 2010; Williams et al., 2011) with a single round of osmium tetroxide staining for thin section TEM (Figures 2C, D and 3B) or electron tomography. In addition, CCM is essentially compatible with a wide range of reactions catalyzed by EM tags other than APEX2 (Ellisman et al., 2015). For example, the protein labeling reactions mediated by miniSOG (Shu et al., 2011) and the tetracysteine-based methods using FIAsH and ReAsH (Gaietta et al., 2002), or the non-protein biomolecule labeling reactions using Click-EM (Ngo et al., 2016) or ChromEM (Ou et al., 2017). The versatility of CCM will likely expand the breath of biological questions that can be addressed using cryofixed samples.

In addition to using EM tags, we have also developed a 3D CLEM protocol (Figure 5A) that allows optimally preserved EM structures to be genetically labeled with fluorescent markers in CCM-processed tissues. In contrast to EM tags, fluorescent markers do not generate electron-dense products (e.g. DAB polymers) that can obscure the subcellular structures. Moreover, with multicolor CLEM, one can utilize multiple readily available genetically encoded fluorescent markers to label different target structures or cells. Using the 3D CLEM protocol, one could also pinpoint labeled subcellular structures (e.g., microtubules) or proteins (e.g., ion channels) in an EM volume with super-resolution microscopy. Furthermore, the ability to genetically label target neurons with fluorescent markers or EM tags in CCM-processed tissues can facilitate circuit reconstructions of identified neurons in optimally preserved specimens.

The advantages of CCM make it particularly suited for addressing biological questions that require optimal and rapid preservation of a genetically labeled structure. For example, to construct an accurate model to describe the biophysical properties of a neuron, it is essential to acquire morphological measurements based on faithfully preserved ultrastructures. CCM processing provides such an opportunity; we were able to obtain a 3D reconstruction of a genetically labeled Drosophila ORN at nanoscale resolution with quality morphological preservation (Figure 3D). In addition, by combining CCM with Flash-and-Freeze EM (Watanabe et al., 2013a, 2013b, 2014a, 2014b) and electron tomography, it is possible to capture the fast morphological changes of genetically labeled vesicles in 3D during synaptic transmission.

Despite its versatility, multiple factors could potentially limit the applicability of CCM. First, given that the core fixation step of CCM is cryofixation, the size of the sample is constrained by the vitrification limit of up to approximately 500 µm (Dahl and Staehelin, 1989; McDonald, 1999; Moor, 1987; Shimoni et al., 1998). In addition, freeze damage due to ice crystal formation can occur (Korogod et al., 2015; Ripper et al., 2008; Shanbhag et al., 2000). Therefore, one should be mindful of freeze damage when performing ultrastructural analysis. Moreover, CCM can only improve the temporal resolution of biological events captured if the specimen is frozen in the live state, but not when the sample was first chemically fixed (e.g. aldehyde-perfused mouse brain). Finally, there are also concerns that some molecules may be lost during rehydration if they are not properly fixed during freeze-substitution (Ripper et al., 2008).

In conclusion, CCM is applicable to addressing questions in diverse tissue types, as demonstrated here with cultured mammalian cells or tissues of Drosophila antennae and mouse brains. Notably, identical solutions and experimental conditions were used for these different tissues in all core steps (Figure 1). Thus, the protocol described here can likely be readily adapted to cells and tissues of other biological systems. In addition, we demonstrated that CCM can further improve the ultrastructure of an aldehyde-perfused brain (Figure 2C and D). Given that aldehyde perfusion is often required for the dissection of deeply embedded or fragile tissues, the compatibility of CCM with aldehyde fixation further broadens the applicability of the method.

A source data file has been provided for Figure 4 (Figure 4-source data 1). The SBEM volume of a Drosophila antenna presented in this study has been deposited to the Cell Image Library. The SBEM volume, the tdTomato confocal volume and the DRAQ5 confocal volume used for 3D CLEM in a mouse brain (corresponding to Figure 5) have also been deposited to the Cell Image Library. The video of 3D CLEM in a mouse brain expressing tdTomato that corresponds to Figure 5-video supplement 1 has been deposited to the Cell Image Library.

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Author details

1. Tin Ki Tsang

   Neurobiology Section, Division of Biological Sciences, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing

   Contributed equally with

   Eric A Bushong

   For correspondence

   t1tsang@ucsd.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1002-106X

2. Eric A Bushong

   National Center for Microscopy and Imaging Research, Center for Research in Biological Systems, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing

   Contributed equally with

   Tin Ki Tsang

   For correspondence

   eric@ncmir.ucsd.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-6195-2433

3. Daniela Boassa

   National Center for Microscopy and Imaging Research, Center for Research in Biological Systems, University of California, San Diego, La Jolla, United States

   Contribution

   Resources, Formal analysis, Funding acquisition, Validation, Investigation, Writing—review and editing

   For correspondence

   dboassa@ncmir.ucsd.edu

   Competing interests

   No competing interests declared

4. Junru Hu

   National Center for Microscopy and Imaging Research, Center for Research in Biological Systems, University of California, San Diego, La Jolla, United States

   Contribution

   Investigation, Writing—review and editing

   For correspondence

   jhu@ncmir.ucsd.edu

   Competing interests

   No competing interests declared

5. Benedetto Romoli

   Department of Psychiatry, School of Medicine, University of California, San Diego, La Jolla, United States

   Contribution

   Resources, Investigation, Writing—review and editing

   For correspondence

   bromoli@ucsd.edu

   Competing interests

   No competing interests declared

6. Sebastien Phan

   National Center for Microscopy and Imaging Research, Center for Research in Biological Systems, University of California, San Diego, La Jolla, United States

   Contribution

   Software, Visualization, Writing—review and editing

   For correspondence

   sph@ncmir.ucsd.edu

   Competing interests

   No competing interests declared

7. Davide Dulcis

   Department of Psychiatry, School of Medicine, University of California, San Diego, La Jolla, United States

   Contribution

   Resources, Funding acquisition, Writing—review and editing

   For correspondence

   ddulcis@ucsd.edu

   Competing interests

   No competing interests declared

8. Chih-Ying Su

   Neurobiology Section, Division of Biological Sciences, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Resources, Supervision, Funding acquisition, Visualization, Writing—original draft, Project administration, Writing—review and editing

   For correspondence

   c8su@ucsd.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-0005-1890

9. Mark H Ellisman

   1. National Center for Microscopy and Imaging Research, Center for Research in Biological Systems, University of California, San Diego, La Jolla, United States
   2. Department of Neurosciences, School of Medicine, University of California, San Diego, La Jolla, United States

   Contribution

   Conceptualization, Resources, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing

   For correspondence

   mark@ncmir.ucsd.edu

   Competing interests

   No competing interests declared

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