The instant application contains a Sequence Listing which has been submitted electronically in ASCII format and is hereby incorporated by reference in its entirety. Said ASCII copy, created on Dec. 17, 2021, is named C1-6182_SL.txt and is 1,984 bytes in size.
The present invention generally relates to methods for preparing live cells for analysis. The invention particularly relates to methods that combine chemical fixation processes with cryofixation processes for localization of membrane proteins.
Localization of membrane proteins via electron microscopy (EM) at high resolution is dependent on robust detection technology and on sample preparation methods that confer superior ultrastructural preservation of membranes. Unfortunately, current methods of localization of membrane-bound proteins at EM resolutions are less than optimal. Immunoelectron microscopy (IEM) to detect either an endogenous or epitope-tagged overexpressed protein using antigenspecific antibodies requires a permeabilization step that also causes degradation of cellular membranes and distortion of membranebound compartments (De Mey et al., 1981; Schnell et al., 2012). An alternative is to fuse enzymatic tags directly to the protein of interest in a transfection experiment, so as to avoid the necessity for introducing an antibody. A number of these enzymatic tags have been described, e.g. metallothioneine (METTEM), resorufin arsenical hairpin (ReAsH), miniSOG, horseradish peroxidase (HRP) and more recently, engineered ascorbate peroxidases (APEX and APEX2) (Hoffmann et al., 2010; Lam et al., 2015; Martell et al., 2012; Mercogliano and DeRoiser, 2007; Porstmann et al., 1985; Shu et al., 2011). While each of these technologies faces its own set of limitations when employed in conjunction with EM, the APEX2 tag, which has been used with success on mitochondrial and ER proteins, is a promising option (Martell et al., 2017). APEX2 is a monomeric 28 kDa soyabean ascorbate peroxidase that withstands strong EM fixation (Lam et al., 2015). Additionally, it is sensitive, generally straightforward in its application and, unlike horseradish peroxidase, is active in both the cytosolic and lumenal compartments (Hopkins et al., 2000; Lam et al., 2015; Martell et al., 2017). Nevertheless, the morphological damage to cellular membranes and membrane-bound organelles that occurs during conventional aldehyde fixation and alcohol dehydration protocols, even without the permeabilization required for antibodies, continues to be an impediment to obtaining optimal preservation of the subcellular architecture.
In contrast, cryofixation or high-pressure freezing (HPF) is a method for obtaining vitreous preparations of live cells and tissues up to 200 μm in thickness with minimal ice crystal formation, thus immobilizing macromolecular assemblies in their near-native state (Chan et al., 1993; Mcdonald, 1999; Zechmann et al., 2007; Studer et al., 2008). This method has become a mainstay for preparing samples for electron tomography, which employs thicker sections (McDonald and Auer, 2006). Further, HPF has been adapted in combination with freeze substitution (FS) methods, which entail organic substitution of water with acetone at low temperature, to generate plastic-embedded samples for conventional EM. However, HPF-FS has not been extensively used with protein localization methods that require chemical fixation of cells (Tsang et al., 2018). Current HPF-FS techniques may not be compatible with common staining processes.
In view of the above, it can be appreciated that there are certain problems, shortcomings or disadvantages associated with localization of membrane proteins via EM, and that it would be desirable if systems and methods were available that were capable of at least partly overcoming or avoiding the morphological damage to cellular membranes and membrane-bound organelles noted above.
The present invention provides methods capable of performing localization of membrane proteins with reduced morphological damage to cellular membranes and membrane-bound organelles relative to conventional techniques.
According to one aspect of the invention, a method is provided for preparing live cells for analysis that includes performing a chemical fixation process on a cellular sample comprising the live cells, and then performing a cryofixation process with extended osmication during freeze substitution on the cellular sample.
According to another aspect of the invention, a method is provided for determining localization of membrane-bound proteins within a preserved membrane architecture within a cellular sample. The method includes performing a glutaraldehyde fixation process on a cellular sample comprising the live cells, performing a cryofixation process with extended osmication during freeze substitution on the cellular sample, performing a staining process on the cellular sample, and analyzing the cell sample with electron tomography.
Technical effects of the methods described above preferably include the capability of fixing the morphology of live cell samples for analysis with little to no damage to cellular membranes and membrane-bound organelles within the sample.
Other aspects and advantages of this invention will be appreciated from the following detailed description.
The intended purpose of the following detailed description of the invention and the phraseology and terminology employed therein is to describe what is shown in the drawings, which include the depiction of one or more nonlimiting embodiments of the invention, and to describe certain but not all aspects of what is depicted in the drawings, including the embodiment(s) depicted in the drawings. The following detailed description also describes certain investigations relating to the embodiment(s) depicted in the drawings, and identifies certain but not all alternatives of the embodiment(s) depicted in the drawings. Therefore, the appended claims, and not the detailed description, are intended to particularly point out subject matter regarded as the invention, including certain but not necessarily all of the aspects and alternatives described in the detailed description.
Disclosed herein are methods for preparing live cells or cellular samples for analysis. The methods include a hybrid approach that combines chemical fixation processes and cryofixation processes. Such methods may be used in combination with staining processes to allow precise localization of membrane proteins in the context of a well-preserved subcellular membrane architecture. Notably, these methods are compatible with electron tomography (EM) and may be used to produce three-dimensional (3D) contextual maps of cellular samples. The methods are broadly applicable to membrane proteins such as luminal and cytosol-facing membrane proteins. Advantageously, the methods may be used to analyze and localize cells grown in monolayers. An exemplary but nonlimiting embodiment includes a method referred to herein as cryoAPEX, which combines glutaraldehyde chemical fixation and high-pressure freezing of cells with peroxidase tagging (APEX). CryoAPEX may include staining with tannic acid and/or uranyl acetate and counter-staining with a lead solution. Optionally, cellular samples may be reacted with diaminobenzadine (DAB). Preferably, the methods do not include certain process steps performed in conventional techniques that cause damage or degradation to the cellular sample, such as but not limited to a dehydration step performed prior to freezing.
Nonlimiting embodiments of the invention will now be described in reference to experimental investigations leading up to the invention.
To evaluate cryoAPEX, an EM tomography-compatible detection method was utilized to visualize the human FIC (filamentation induced by cAMP) protein, HYPE (also known as FICD). FIC proteins are a recently characterized class of enzymes that predominantly utilize ATP to attach AMP (adenosine monophosphate) to their protein targets (Casey and Orth, 2018; Truttmann et al., 2017; Worby et al., 2009). This post-translational modification is called adenylylation or AMPylation. The first FIC proteins, VopS and IbpA, were described in the pathogenic bacteria Vibrio parahemolyticus and Histophilus somni, respectively, where they serve as secreted bacterial effectors that induce toxicity in host cells by inactivating small GTPases through AMPylation (Mattoo et al., 2011; Worby et al., 2009; Xiao et al., 2010; Yarbrough et al., 2009; Zekarias et al., 2010). FIC proteins have also been implicated in bacterial cell division and persister cell formation, protein translation, cellular trafficking and neurodegeneration (Garcia-Pino et al., 2014; Harms et al., 2015; Mukherjee et al., 2011; Truttmann et al., 2018).
HYPE (huntingtin yeast interacting protein E) or FICD is the sole FIC protein encoded by the human genome (Faber et al., 1998; Sanyal et al., 2015). In humans, HYPE is expressed ubiquitously, albeit at very low levels. It is a single-pass type II membrane protein that localizes to the lumenal surface of the endoplasmic reticulum (ER) (Sanyal et al., 2015; Worby et al., 2009). HYPE plays a critical role in regulating ER homeostasis by reversibly AMPylating the Hsp70 chaperone, BiP (also known as HSPA5) (Ham et al., 2014; Sanyal et al., 2015; Preissler et al., 2015, 2017a,b). Biochemical and proteomic screens have identified additional AMPylation targets of HYPE and its orthologs, which include cytosolic chaperones, cytoskeletal proteins, transcriptional and translational regulators, and histones (Broncel et al., 2016; Truttmann et al., 2016, 2017). These data suggest that a fraction of HYPE could reside outside the ER, for example, in the nucleus or cytoplasm. Indeed, a small fraction of the HYPE homolog in Caenorhabditis elegans, FIC-1, has been shown to localize to the cytosol and AMPylate cytosolic c and Hsp40 proteins (Truttmann et al., 2017).
The low levels of HYPE expression in human cells combined with the resolution limitations of conventional immunofluorescence microscopy make obtaining definitive localization data difficult. CryoAPEX circumvents these limitations by using an electron microscopy approach. To visualize HYPE in cells, a technique was developed that preserves membrane ultrastructure and is compatible with transmission EM tomography methods to identify specific distribution of HYPE in three-dimensional (3D) space. HYPE was visualized by genetically tagging it with APEX2. The APEX2 tag catalyzes a peroxide-based reaction that converts diaminobenzadine (DAB) into a low-diffusing precipitate that deposits at the site of the target protein (Lam et al., 2015). In an analysis of HYPE leading to certain aspects of the present invention, a peptide, designated endoplasmic reticulum membrane (ERM), was provided to serve as a dual control for ER localization as well as for ER morphology. ERM consists of the N-terminal 1-27 amino acids of cytochrome P450 2C1 (CYP2C1) (Lam et al., 2015; Sandig et al., 1999). An ERM-APEX2 fusion localizes to the ER membrane such that the APEX2 tag faces the cytosol. Additionally, ERM is known to induce a reorganization of the smooth ER into distinctive ordered membrane structures called organized smooth ER (OSER) (Snapp et al., 2003; Lam et al., 2015; Sandig et al., 1999). Thus, ERM-APEX2 serves as an excellent metric for assessing both ER membrane-specific staining and ultrastructural membrane preservation.
Next, since degradation of the cellular ultrastructure in traditional aldehyde fixation and alcohol dehydration methods appears to be largely associated with the alcohol dehydration post-processing steps and not with aldehyde fixation per se, a combination method proposed by Sosinsky et al. (2008) was utilized, which relies on chemical fixation prior to cryofixation and optimally preserves membrane structure. This combination approach was applied to the detection of APEX2-tagged proteins. Using this methodology, minimal lipid extraction or distortion of membrane structures was observed, and were able to clearly detect ER membrane-bound HYPE.
The hybrid method (cryoAPEX) performed remarkably well for protein localization at the subcellular level. Comparison of ERM-APEX2 in cryoAPEX-treated cells versus live cryofixed (HPF alone) cells showed well-preserved OSER morphology with ER-specific staining only on the cytosolic face of the ER membrane. Further, data collected on cryoAPEX-treated cells could be used to reconstruct a 3D representation of HYPE within the ER lumen. This ability to simultaneously assess the detection of a membrane protein in multiple cellular compartments throughout the subcellular volume of a single cell at low-nanoscale resolution is a significant advance. More broadly, cryoAPEX presents a straightforward methodology for probing the subcellular distribution of membrane-bound proteins, with either lumen-facing or cytosol-facing topologies, that is amenable to high-resolution 3D tomographic reconstruction.
In these investigations, HYPE-APEX2 plasmids were provided and analyzed. HYPE-APEX2 fusion was first synthesized and then inserted into pcDNA3 vector between BamHI and XhoI sites. Briefly, a fusion construct comprising the first 118 amino acids of the mouse isoform of α-mannosidase with the APEX2 gene in its C-terminus following a short intervening linker sequence was first synthesized and then cloned into pcDNA3.1. The complete sequence for MannII-APEX2 is provided (Table S1 in
Transfection and chemical fixation were performed by growing HEK-293T cells (ATCC) in 10 cm dishes in Dulbecco's modified Eagle's medium (DMEM; Corning) supplemented with 10% fetal bovine serum (FBS; Corning NuSerum IV) at 37° C. and 5% CO2. Cells were transfected with APEX2-tagged mammalian expression plasmids using Lipofectamine 3000 (Thermo Fisher). Cells were washed off the plate with Dulbecco's PBS 12 to 15 h post-transfection and then pelleted at 500 g. For those samples requiring chemical fixation, pellets were resuspended in 0.1% sodium cacodylate buffer containing 2% glutaraldehyde for 30 min, washed 3 times for 5 min with 0.1% sodium cacodylate buffer and 1 time with cacodylate buffer containing 1 mg/ml 3,3′-diaminobenzidine (DAB) (Sigma-Aldrich). Pellets were then incubated for 30 min in a freshly made solution of 1 mg/ml DAB and 5.88 mM hydrogen peroxide in cacodylate buffer, pelleted and washed 2 times for 5 min in cacodylate buffer and once with DMEM. Finally, cell pellets were resuspended in DMEM containing 10% FBS and 15-20% BSA, then pelleted again. The supernatant was aspirated and excess media wicked off with a Kimwipe in order to remove as much liquid as possible. For BHK (ATCC) controls, cells were grown in DMEM with 10% FBS at 37° C. and 5% CO2. Cells were either cryofixed directly or prefixed with glutaraldehyde prior to cryofixation. An identical freeze-substitution protocol was used for processing for both HEK-293T and BHK cells. BHK cells were post-stained with uranyl acetate and Sato's lead prior to imaging.
To perform high-pressure freezing and freeze substitution, cell pellets (2 to 3 μl) were loaded onto copper membrane carriers (1 mm×0.5 mm; Ted Pella Inc.) and cryofixed using the EM PACT2 high-pressure freezer (Leica). Cryofixed cells were then processed by freeze substitution using an AFS2 automated freeze substitution unit (Leica). An extended freeze substitution protocol was optimized for the preferential osmication of the peroxidase-DAB byproduct. Briefly, frozen pellets were incubated for 24 h at −90° C. in acetone containing 0.2% tannic acid and then washed 3 times for 5 min with glass-distilled acetone (EM Sciences). Pellets were resuspended in acetone containing 5% water and 1% osmium tetroxide, with or without 2% uranyl acetate (as applicable) for 72 h at −80° C. Following this extended osmication cycle, pellets were warmed to 0° C. over 12 to 18 h. Pellets were then washed 3 times for 30 min with freshly opened glass-distilled acetone. Resin exchange was carried out by infiltrating the sample with a gradually increasing concentration of Durcupan ACM resin (Sigma-Aldrich) as follows: 2%, 4% and 8% for 2 h each and then 15%, 30%, 60%, 90%, 100% and 100% plus component C (Durcupan100+C hereafter) for 4 h each. Resin-infiltrated samples in membrane carriers were then embedded in resin blocks and polymerized at 60° C. for 36 h. Post-hardening, planchets were extracted by dabbing liquid nitrogen on the membrane carriers and using a razor to resect them out of the hardened resin. After extraction of membrane carriers, a thin layer of Durcupan100+C was added on top of the exposed samples and incubated in an oven at 60° C. for 24 to 36 h to obtain the final hardened sample blocks for sectioning.
Sample preparation was performed via a conventional room temperature method. Specifically, HEK-293T cells were grown on collagen-coated glass coverslips and transfected with HYPE-APEX2 or ERM-APEX2 mammalian expression plasmids for 24 h as above. Cells were then washed with DPBS and chemically fixed with 2% glutaraldehyde in 0.1% sodium cacodylate buffer for 30 min. Fixed cells were washed with cacodylate buffer and finally with 1 mg/ml of DAB in cacodylate buffer for 2 min. Following the wash, cells were incubated for 30 min in a freshly made solution of 1 mg/ml of DAB and 5.88 mM of hydrogen peroxide in cacodylate buffer at room temperature. Cells were washed 3 times for 2 min each with DPBS, incubated in an aqueous solution of 1% osmium tetroxide for 10-15 min and then washed with distilled water. Dehydration was carried out using increasing concentration of 200 proof ethanol (30%, 50%, 70%, 90%, 95%, 100%) followed by resin infiltration of the cells with gradually increasing concentrations of Durcupan resin in ethanol (30%, 60%, 90%, 100%), then Durcupan100+C. Coverslips were placed on BEEM capsules filled with Durcupan100+C, cell-face-down on the resin and incubated in an oven for 48 h at 60° C. After polymerization, coverslips were extracted by dipping the coverslip face of the blocks briefly in liquid nitrogen. Serial sections were then obtained by sectioning of the blocks en face and ribbons collected on formvar-coated slot grids.
For serial sectioning, lead staining and electron microcopy, thin (90 nm) serial sections were obtained using a UC7 ultramicrotome (Leica) and collected onto formvar-coated copper slot grids (EMsciences). Glass knives were freshly prepared from glass sticks during each sectioning exercise. Lead staining of the sections was carried out for 1 min wherever applicable with freshly made Sato's lead solution. Samples were screened on a Technai T-12 80 kV transmission electron microscope, and an average of 15-20 cells from multiple blocks were visualized for each sample.
EM tomography, data reconstruction, and segmentation included using thicker (250 nm) sections for collecting tomographic tilt-series. Sections were coated with gold fiducials measuring 20 nm in diameter prior to collection. Tilt-series of a single 250 nm section were collected with automation using the program SerialEM (Mastronarde, 2003) on a JEOL3200 TEM operating at 300 kV. The collected tilt-series were then aligned and tomogram generated by weighted back projection using the eTomointerface of IMOD (Kremer et al., 1996). The reconstructed tomogram was visualized in IMOD. The ER membrane was first hand-segmented and then used as a mask for thresholding of the density within the ER lumen.
In cellular imaging, the ability to obtain 3D spatial localization of proteins in the context of well-preserved cellular structures at high resolution is highly desirable (O'Toole et al., 2018). For antibody-conjugate-based detection methods to be effective, ultrastructure-damaging permeabilization and/or technically challenging ultracryosectioning are required. At EM resolutions, the deleterious effects of such treatments, particularly upon membrane-bound compartments, become obvious. Fusion of a protein of interest to the monomeric enhanced peroxidase APEX2 avoids the need for an antibody. However, the published protocols still use chemical fixation and alcohol dehydration prior to embedding (Martell et al., 2017).
It was hypothesized that the alcohol dehydration step in published APEX2 protocols is limiting for ultrastructural preservation, and consequently for signal intensity and resolution. Therefore, cryofixation-freeze substitution (HPF-FS) of live cells was utilized for preservation, and it was combined with chemical fixation methods to optimize ultrastructural preservation in untransfected human HEK-293T cells.
Preservation was assessed based on several criteria including membrane integrity, smoothness of intracellular membranes, densely packed cytoplasm, and maintenance of organellar structures such as mitochondria with clearly visible cristae. Smoothness of intracellular membranes is often a primary indication of good preservation, and preservation of the nuclear envelope has been used classically as a hallmark (Sosinsky et al., 2008; Tsang et al., 2018). As shown in
Next, the applicability of the combination method was tested for ultrastructural preservation with the use of the APEX2 tag to follow a protein of interest, creating the hybrid glutaraldehyde+HPF-FS+APEX2 method referred to herein as cryoAPEX (
To serve as a proof of concept, cryoAPEX was evaluated by assessing the membrane localization of the ERM-APEX2 chimeric protein (
As a morphological control, thin sections of ERM-APEX2-transfected cells were processed using HPF-FS alone and then stained using osmium tetroxide, tannic acid and uranyl acetate but without chemical fixation or the DAB reaction. Under these conditions, the typical membrane whorl pattern of reorganized ER adjacent to the nucleus was clearly visible (
To enhance the contrast of membrane staining over the background of overall osmicated DAB density (i.e. osmiumstained DAB precipitate), the same sections shown in
Having established the methodology to localize an APEX2-tagged ER membrane protein in an optimally preserved cell, cryoAPEX was next applied to the sole human FIC protein, HYPE. Previous immunofluorescence, cell fractionation and protease protection assays have placed HYPE as predominantly facing the ER lumen (Sanyal et al., 2015). A smaller fraction of HYPE has also been detected in the cytosol, as well as in the perinuclear space (Truttmann et al., 2015, 2017). Therefore, HEK-293T cells were transiently transfected with HYPE-APEX2 constructs and processed using cryoAPEX as standardized for ERM-APEX2 above. As before, ultrastructure was well preserved and there were minimal signs of lipid extraction or membrane ruffling (
Further, this density was not an artifact of the rough ER, where such density corresponding to the presence of ribosomes is seen only on the cytosolic face of the ER membrane (
Perhaps the greatest advantage of using cryofixation for sample preparation for EM is its capability to preserve membrane ultrastructure consistently throughout the cell volume. To track the distribution of HYPE over a large subcellular volume, serial section EM was employed (
To visualize HYPE, a tilt series from a single 250 nm section of HEK-293T cells expressing HYPE-APEX2 was collected (
Next, 3D modeling by segmentation and visualization of the HYPE-APEX2 density showed that HYPE was confined within the ER membrane (
FIC-mediated adenylylation (AMPylation) is an important, evolutionarily conserved mechanism of signal transduction. In humans, AMPylation mediated by HYPE regulates the unfolded protein response via reversible modification of BiP (Ham et al., 2014; Preissler, et al., 2017a,b; Sanyal et al., 2015). It is an open question as to whether HYPE functions beyond its role as a UPR regulator with additional physiological targets. Indeed, despite the fact that it has previously been shown that HYPE is an ER membrane protein that faces the lumen (Sanyal et al., 2015; Rahman et al., 2012), a number of candidate targets have recently emerged for further evaluation that reside outside the ER (Broncel et al., 2016; Truttmann et al., 2016). For instance, the C. elegans homolog of HYPE (FIC-1) can be detected in the cytosol and is implicated in controlling the function of cytosolic chaperones (Truttmann et al., 2017). The Drosophila melanogaster HYPE homolog (dFic) is implicated in blindness and has been shown to associate with cell surface neuro-glial junctions, possibly by entering the secretory pathway (Rahman et al., 2012). Thus, a clear subcellular localization for HYPE is needed to better understand its role in the context of these new protein targets—which led to the development a technique to determine the subcellular localization of membrane proteins like HYPE at a high (low-nanoscale) resolution.
Despite tremendous advances in light microscopy, electron microscopy still remains the technique of choice to visualize cellular ultrastructure or determine protein localization at nanoscale resolution. Here, the development of an EM method, called cryoAPEX, is described which successfully adapts the APEX2 tag for cryofixation while simultaneously retaining membrane preservation. Additionally, it has been evidenced herein that data obtained using cryoAPEX for visualizing ER proteins, HYPE and ERM, can be used for EM tomographic reconstruction of membranes in 3D. Applying cryoAPEX to HYPE localization, it has been shown herein that HYPE appears to reside solely in the ER lumen and in the contiguous nuclear envelope, in agreement with immunofluorescence data from industry (Sanyal et al., 2015; Truttmann et al., 2015).
CryoAPEX is designed specifically for localizing membrane-bound proteins. The methodology presented herein enables sufficient resolution and membrane preservation such that even structures in close contact with the ER, such as the Golgi, mitochondria or plasma membrane, are clearly distinguishable as HYPE-negative (
Many organelles and transport vesicles within a cell are labile structures that are difficult to preserve in their native morphology. An organelle like the ER has multiple domains that make contacts with several other organelles as well as the plasma membrane. The functional relevance of these organellar contact points is of research interest and, in the case of the ER, they are known to be portals of lipid and calcium transport (English and Voeltz, 2013; Rowland and Voeltz, 2012). Thus, preservation of these structures was especially important for ascertaining the distribution of HYPE.
Next, the applicability of various traditional protein localization techniques were considered such as immunoelectron microscopy (IEM), metal-tagging EM (METTEM), and peroxidase tagging. Unfortunately, each of these techniques suffers from a variety of limitations in addition to inadequate sample preservation. Specifically, current methods of detection are based on two common processes: (1) a chemical fixation step that precedes the actual detection assay and (2) a sample preparation step involving dehydration of fixed cells via alcohol at room temperature or on ice. This combination of chemical reagents leads to poor preservation of the membrane morphology as a result of lipid extraction, and introduces artefacts. Therefore, in addition to membrane preservation, the method of choice needs to be compatible with heavy metal staining, so as to impart an adequate level of contrast between HYPE-associated membranes and other organellar membranes for contextual information about the ultrastructural environment within which HYPE resides. This ruled out METTEM tagging, as the technique is incompatible with the use of heavy metal stains (Risco et al., 2012).
Lastly, the amenability of the method to 3D electron microscopic techniques was considered. This is notable as organelles or membrane structures such as the ER, Golgi, mitochondria, or the plasma membrane cover a vast three-dimensional subcellular space and are in a constant state of morphological equilibrium with their surroundings. They undergo constant remodeling in their different domains in response to functional cues that can alter the localization of proteins that are associated with them (Shibata et al., 2010; Voeltz et al., 2002). To detect such changes or, alternatively, the exclusive localization of a target protein in specific domains of these large organelles, 3D information at the site of protein localization can yield critical clues about protein function. Thus, development focused on a method that incorporated each of the above criteria to yield HYPE's subcellular distribution in an optimally preserved and 3D EM-compatible sample.
Cryofixation of live cells under high pressure (HPF) is a method that shows improved ultrastructural preservation and is now routinely used to prepare samples for EM tomography (McDonald and Auer, 2006; O'Toole et al., 2018). It is not deemed compatible with most of the detection methods described above, however, as they require chemical fixation. Thus, to determine the subcellular distribution of HYPE, cryoAPEX was developed, a hybrid method that combines the power of APEX2 genetic tagging and HPF cryofixation. Chemically fixed cells expressing APEX2-tagged HYPE were first reacted with DAB to generate HYPE-specific density, and then cryofixed and freeze-substituted with acetone. As shown, cryoAPEX not only displays specificity of detection at high resolution for both lumen-facing (HYPE) and cytosol-facing (ERM) ER membrane proteins, but also retains ultrastructural preservation that makes cryoAPEX amenable to TEM tomography. Further, cryoAPEX can be used to assess cells grown in monolayers, making it widely applicable.
A notable aspect of cryoAPEX is the robustness of the DAB byproduct that can withstand a long freeze-substitution reaction in acetone. It was observed that once chemically fixed and labeled with DAB, cells do not need to be cryofixed right away. For example, aldehyde-fixed, DAB-labeled cells that were cryofixed after 48 h (storage at 4° C.) exhibited no deterioration of cellular ultrastructure and staining when compared to those that were cryofixed immediately. This feature could prove to be of great advantage to laboratories that do not have immediate access to HPF and freeze substitution units.
In conclusion, cryoAPEX has been shown to be a method for obtaining localization of a single APEX2-tagged protein at a high resolution while maintaining excellent ultrastructural preservation and compatibility with EM tomography. CryoAPEX was applied to assess the subcellular localization of the human FIC protein, HYPE, and show that it is robustly detected very specifically on the lumenal face of the ER membrane and in cellular compartments that are contiguous with the ER lumen, where it displays periodic distribution resembling possible signaling complexes. Further, HYPE was not detected in the mitochondria, nucleus, plasma membrane, or the Golgi and secretory network at the expression levels tested. Additionally, it was shown that cryoAPEX works equally well for cytosol-facing membrane proteins, such as ERM, and accurately reflects ultrastructural morphological changes. Finally, it was demonstrated that cryoAPEX can be applied to assessing protein localization using cell monolayers and executed in basic cell biology laboratories with relative ease.
While the invention has been described in terms of specific or particular embodiments and investigations, it should be apparent that alternatives could be adopted by one skilled in the art. For example, process parameters such as temperatures and durations could be modified, and appropriate materials could be substituted for those noted. Accordingly, it should be understood that the invention is not necessarily limited to any embodiment described herein. It should also be understood that the phraseology and terminology employed above are for the purpose of describing the disclosed embodiments and investigations, and do not necessarily serve as limitations to the scope of the invention. Therefore, the scope of the invention is to be limited only by the following claims.
This application claims the benefit of U.S. Provisional Application No. 63/081,105, filed Sep. 21, 2020, the contents of which are incorporated herein by reference.
This invention was made with government support under Contract Nos. AI081077 and GM100092 awarded by the National Institute of Health. The government has certain rights in the invention.
Number | Date | Country | |
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63081105 | Sep 2020 | US |