Ultrastructural visualization of trans-ciliary rhodopsin cargoes in mammalian rods
© Chuang et al.; licensee BioMed Central. 2015
Received: 1 October 2014
Accepted: 15 January 2015
Published: 8 February 2015
Cilia are vital to various cellular and sensory functions. The pathway by which ciliary membrane proteins translocate through the transition zone is not well understood. Direct morphological characterization of ciliary cargoes in transit remains lacking. In the vertebrate photoreceptor, rhodopsin is synthesized and transported from the inner segment to the disc membranes of the outer segment (OS), which is a modified cilium. To date, the membrane topology of the basal OS and the mechanisms by which rhodopsin is transported through the transition zone (i.e., connecting cilium) and by which nascent disc membranes are formed remain controversial.
Using an antibody recognizing its cytoplasmic C-terminus, we localize rhodopsin on both the plasma membrane and lumen of the connecting cilium by immuno-electron microscopy (EM). We also use transmission EM to visualize the electron-dense enzymatic products derived from the rhodopsin-horseradish peroxidase (HRP) fusion in transfected rodent rods. In the connecting cilium, rhodopsin is not only expressed in the plasma membrane but also in the lumen on two types of membranous carriers, long smooth tubules and small, coated, filament-bound vesicles. Additionally, membrane-bound rhodopsin carriers are also found in close proximity to the nascent discs at the basal OS axoneme and in the distal inner segment. This topology-indicative HRP-rhodopsin reporter shows that the nascent basalmost discs and the mature discs have the same membrane topology, with no indication of evagination or invagination from the basal OS plasma membranes. Serial block face and focus ion beam scanning EM analyses both indicate that the transport carriers enter the connecting cilium lumen from either the basal body lumen or cytoplasmic space between the axonemal microtubules and the ciliary plasma membrane.
Our results suggest the existence of multiple ciliary gate entry pathways in rod photoreceptors. Rhodopsin is likely transported across the connecting cilium on the plasma membrane and through the lumens on two types of tubulovesicular carriers produced in the inner segment. Our findings agree with a previous model that rhodopsin carriers derived from the cell body may fuse directly onto nascent discs as they grow and mature.
The cilium is a vital organelle harboring receptors and channels for a variety of sensory functions. Human mutations of genes important for ciliary structure and/or protein trafficking have been linked with ciliopathic diseases. The pathway by which ciliary membrane proteins translocate from the cell body through the proximal diffusion barrier, known as the “transition zone (TZ),” remains to be elucidated. Prior studies have used combinations of biochemistry, reverse genetics, and functional analyses, but morphological delineation of ciliary membrane cargoes during TZ transit has been largely lacking.
The TEM images of rodent rods obtained in our lab did not show evidence of outwardly folded PM between the CC and the OS . The few basalmost nascent discs/cisternae are smaller than the mature discs (Figure 2C), morphologically resembling the evaginated PM described by Steinberg et al.  and others [15,16], except that they are enveloped by the OS PM. More recently, using cryo-EM, a method without chemical fixation, Gilliam et al.  showed that the base of the rod OS is completely “closed” by the PM as well. Due to the low tissue penetration of cryo-EM method, dissociated rod axonemes were used, so concerns remain whether the extensive axoneme isolation procedures may have had an impact on the OS membrane structure.
Primarily based on rhodopsin immunolocalization studies [1,8,18-24], several models have been proposed to explain the transport route of rhodopsin from the inner segment to the OS. The current predominant model proposes that rhodopsin translocates through the CC along the ciliary PM via intraflagellar-mediated transport (IFT) [25-28]. In agreement with this model, post-embedding immuno-EM detected rhodopsin on the CC PM [23,29,30]. Furthermore, transgenic mice constitutively null for KIF3A (a key motor that drives IFT)  and hypomorphic for IFT88 (a key IFT component)  had rhodopsin mislocalized from the OS. Later studies using conditional knockout mice in which kif3A was deleted after OS maturation, however, suggested that KIF3A is dispensable for the OS localization of rhodopsin . Thus, whether or not the IFT pathway is critically involved in rhodopsin ciliary transport remains unclear.
Vesicular structures have been previously seen in the CC lumen of both developing and adult rods through the use of the rapid freeze/etch, TEM, and cryo-EM methods [10,14,17]. Additionally, several disease mouse models exhibit membrane-bound vesicles abnormally accumulated at the base of the rod CC, in the CC lumens, the basal OS axonemes, and/or the extracellular space near the CC; this phenotype has often been associated with rhodopsin mislocalization [14,17,32,33]. While indicative, direct evidence demonstrating that the CC vesicles are the carriers for rhodopsin remains lacking.
Previous studies showed that horseradish peroxidase (HRP), when being expressed at the luminal side of a membrane protein, can be used as a genetic tag for ultrastructural localization study based on the electron-dense enzymatic products derived from HRP activity [34-36]. HRP expression causes little or no effect on membrane integrity, and its relatively uniform labeling allows identification of small intracellular structures, such as synaptic vesicles . Unlike many antigen epitopes, HRP activity is less affected by microenvironmental pH and glutaraldehyde fixation . Thus, HRP can be visualized in hard-fixed samples with better-preserved ultrastructural morphology. Furthermore, the HRP substrates, H2O2 and 3,3′-diaminobenzidine (DAB) tetrahydrochloride hydrate, are far smaller than a nano-gold-conjugated antibody (Ab) and therefore have better tissue penetration. This is especially important when detergent is omitted during the labeling procedure to better preserve membrane ultrastructure.
In this paper, we have combined immuno-EM, TEM, and three-dimensional scanning EM (3D-SEM) approaches to comprehensively delineate the nature of rhodopsin-bearing transport carriers and their spatial relationship to the CC, as well as the membrane topology of the basal OS. These findings greatly improve our understanding of disc genesis and rhodopsin’s OS transport pathways.
Reagents and animals
A cDNA fragment encoding a signal sequence fused to HRP was PCR amplified from the ssHRP-TM vector  and inserted into the N-terminus of rhodopsin-GFP-1D4 plasmid (1D4 is an epitope encoded by ETSQVAPA) . The addition of the signal sequence and resulting glycosylation of HRP are necessary for its enzymatic activity . The entire coding sequence of ssHRP-rhodopsin-GFP-1D4 plasmid was then transferred to the pCAG vector (gift of Connie Cepko ) for electroporation. All chemicals were purchased from Sigma unless otherwise mentioned. Rabbit anti-rhodopsin C-terminus Ab C107 was generated using maltose-binding protein fusion containing rhodopsin’s C-terminal 39 residues as an antigen (Cocalico Biochemicals, PA). A 10-nm gold-conjugated anti-rabbit IgG (Electron Microscopy Sciences) was also used. Isolation of bovine rod OS  and Odyssey-based immunoblotting assay were carried out using standard protocols.
All methods that involved live animals were approved by the Weill Medical College of Cornell University Institutional Animal Care and Use Committee.
For post-embedding immuno-EM, CD1 mouse retinas were harvested by transcardial perfusion with 20 mL of 4% paraformaldehyde plus 0.1% glutaraldehyde. Retina blocks were quenched in 0.1 M glycine in phosphate-buffered saline (PBS) for 5 min, rinsed with PBS, and followed by gradient dehydration in ethanol solutions. Retina blocks were then embedded in LR-White resin (Electron Microscopy Sciences) and polymerized for 48 h at 50°C, following the manufacturer’s instructions. Ultrathin sections (70 nm) were cut and collected on nickel grids. The grids were then baked at 60°C for 1 h. Prior to immunolabeling, grids were etched with 1% H2O2 and rinsed in deionized water. Grids were blocked by 1% bovine serum albumin (BSA)/PBS and then incubated with primary Abs at room temperature overnight. Grids were washed in PBS and incubated with 10-nm gold-conjugated secondary Abs (Electron Microscopy Sciences) in PBS containing 1% BSA/fish gelatin (Amersham Biosciences) at room temperature. After final washes in PBS and then water, grids were counter stained with uranyl acetate and lead citrate for final examination on a Philips CM10 electron microscope. Approximately 150 rods were examined on the scope, and electron micrographs of a total of 52 rods were taken for detailed analysis.
Retinal transfection, light microscopy examination, HRP activity detection, and TEM
For rod transfection, 2 μg of HRP-rhodopsin reporter plasmid was injected subretinally, followed by electroporation of the eyes of postnatal day 0 Sprague-Dawley rats, as described [39,41]. Animals were housed in 12-h light-dark cycles up to postnatal day 21 (at which time the rods had reached maturation) and harvested around noon under normal room light using cardiac perfusion, as previously described . Retinal pieces (~2 × 4 mm) isolated from fixed eyecups were embedded in 5% low-melting agarose and cut by vibratome (40 μm thick). Green fluorescence in transfected rods was directly observed by Leica TCS SP2 spectral confocal microscopy.
For HPR detection, freshly cut, fixed retina vibrotome sections were quenched in 50 mM NH4Cl/0.1 M phosphate buffer, pH 7.4 (PB) for 10 min or 1% NaBH4/PB for 30 min at room temperature. After washes in Tris-buffered saline, sections were treated with 0.22 mg/mL DAB in Tris-buffered saline plus 1.5% H2O2 and monitored under a microscope for proper development of chromogenic products, typically 4–12 min. The reaction time was chosen with the aim of developing the DAB precipitates for a good signal-to-noise contrast, rather than the highest labeling density. Reactions were stopped by transferring sections to fresh Tris-buffered saline and then to PB. Sections were then subjected to EM analysis following routine procedures . Briefly, the labeled vibrotome sections were fixed by 2% osmium tetroxide in PB, dehydrated with graded ethanol, embedded in Epon, and then subjected to ultrathin sectioning. The ultrathin sections were then treated with uranyl acetate-lead citrate counterstaining before examination under a Philips CM10 microscope.
Serial block face scanning electron microscopy
Adult CD1 mice were transcardially perfused with a mixture of 2.5% glutaraldehyde and 4% paraformaldehyde in 0.1 M cacodylate buffer, pH 7.4. The eyes were then enucleated and post-fixed for an additional 2 days on ice in the same solution. Small pieces of eyecup (~2 × ~2 mm) were processed for en bloc fixation and staining, as described [42,43]. Briefly, specimens were incubated with 1.5% potassium ferrocyanide, 2 mM calcium chloride, and 2% osmium tetraoxide in 0.15 M cacodylate buffer, pH 7.4 for 1 h on ice followed by treatment with thiocarbohydrazide solution for 20 min at room temperature then with 2% osmium tetroxide fixation for 30 min at room temperature. After fixation, the tissue block was incubated in 1% uranyl acetate at 4°C overnight. The following day, en bloc Walton’s lead aspartate staining was performed at 60°C for 30 min, followed by serial ethanol dehydration and embedding in Embed-812 (Electron Microscopy Sciences). Tissue blocks were mounted and electrically grounded with the application of silver paint. The surface of the specimen was sputter coated with a thin layer of gold/palladium and subjected to serial block face imaging using a 3View ultramicrotome. Samples were imaged every 65 nm at a resolution of ~3.5 nm per raw pixel.
Focused ion beam scanning electron microscopy
Retinal tissue blocks of adult C57BL/6J mice were prepared and en bloc stained as described above. Block face images of the samples that were precision milled every 10 nm (with a pixel resolution of ~5 nm/pixel in x, y, and z) or 5 nm (with a pixel resolution of ~3.5 nm/pixel in x, y, and z) were collected by FEI Helios NanoLab 650 microscope. Images were processed using Serial Sections Alignment Programs of IMOD/eTomo to correct drifting caused by the 30° angle from the block face during imaging.
Results and discussion
Immuno-EM of rhodopsin in mouse rod CC
Immunolabeling patterns can be influenced by several factors (e.g., availability of the antigen epitopes, fixation/embedding/permealization conditions). Previous studies by Hicks and Molday  found that, under the same labeling conditions, rhodopsin was predominantly labeled on the plasma membranes by an Ab against the extracellular N-terminus but predominantly labeled on the disc membranes by an Ab against the cytoplasmic C-terminus. They explained the difference as due, in part, to the accessibility of the antigen epitope.
The CC immunolabeling pattern of rhodopsin could also be influenced by the Ab and/or the conditions used. A previous study demonstrating the CC PM location of rhodopsin was carried out using a monoclonal Ab recognizing its N-terminus (facing the extracellular side) . We decided to revisit the CC location of rhodopsin by performing immuno-EM of rhodopsin in mouse rods using a polyclonal Ab specifically recognizing rhodopsin’s C-terminus (Figure 1D; also see “Methods”). As predicted , the OS discs were heavily labeled by rhodopsin immunogolds. Rhodopsin immunogolds were also detected in the distal inner segments near basal bodies as well as in the CC (white arrows, Figure 1D). In the CC, rhodopsin immunogolds were localized to both the PM and the lumen in a roughly 1:2 ratio (n = 52). These labelings were immunospecific; they were absent when the primary Ab was omitted or when the primary Ab was presorbed by the antigen (not shown).
While these studies clearly indicated that rhodopsin is expressed in the lumen of the CC, the limited membrane preservation offered by the immuno-EM prevented us from characterizing the types of the transport carriers nor the detailed location of the rhodopsin distributed within the CC lumen.
Characterization of the membrane topology of the OS base
Expression studies of ectopically expressed reporter molecules have been recognized as a useful means to complement the immunolabeling of endogenous molecules and to circumvent the technical problems associated with immunolabeling. Previous studies have shown that rhodopsin reporters transfected in rodents rods are predominantly localized to the OS [14,41]. Several retinitis pigmentosa mutant rhodopsins share a similar mislocalization pattern in transfected rat rods and conventional transgenic mouse rods . These results indicate that transfected rhodopsin follows the same transport pathway used by endogenous rhodopsin. Thus, we set out to visualize the ultrastructural distribution of HRP-tagged rhodopsin reporter in transfected rods in situ based on electron-dense DAB reaction products. Since we tagged the HRP onto the N-terminus of rhodopsin (Figure 2A), the DAB precipitates are expected on the extracellular side of the PM, the luminal side of the disc membrane, and the luminal side of rhodopsin transport carriers. We reasoned that the precise distribution of the DAB deposits, as well as their interrelationship with the surrounding cytosol, could unambiguously demonstrate the membrane topology of the basal OS membranes (Figure 2B vs. 2C).
In either non-transfected or transfected rods, the PM and the cytosol of the OS surrounded the entire disc stacks, including the most proximal cisternae (arrowheads, Figure 3A) and tubulovesicles (black arrows, Figure 3B). The previously described “open discs” at the basal OS with their extracellular surface labeled by DAB precipitates were not detected. No internalized membrane profiles indicative of invagination were observed either.
Others have considered the possibility that the extra PM artificially gained during the chemical fixation (or other experimental procedures) might somehow encompass the evaginated outfolds, rendering a “closed” appearance to the OS base [2,8]. If this were true, one would expect to see the DAB products on the convex sides of these “folds” (Figure 2B), regardless of whether or not they are enwrapped by the PM. That was however not the case. All of the basalmost discs had the DAB products expressed inside their lumens (Figure 3B, C).
Ultrastructural characterization of the rod CC axoneme
The second type of structures had a particle appearance. However, close inspection showed that these structures were, in fact, small vesicles. These vesicles had heavy coats, and an estimated diameter was up to ~20 nm (arrowheads, Figure 4A; black arrows, Figure 4E–G). These vesicles were distributed either singly or tethered to each other by thin “tuft-like” filaments (Figure 4A, E–G). These vesicles were predominantly found in inner shafts (Figure 4A, E–G) and to a lesser degree in the outer shaft (Figure 4A, F). In some cases, the coated vesicles were tightly clustered, rendering a dark fuzzy ball-like structure (black curved arrows, Figures 4F and 5A–D). The vesicle clusters varied in both size (~60–100 nm in diameter) and shape and were found anywhere between the proximal CC to the basal OS axonemes (but rarely in the distal OS axoneme). The heterogeneous expression pattern of the dark fuzzy structures suggests they are not the counterpart of the amorphous electron-dense matrix structure seen in the basal body lumens of motile cilia [48,49].
Morphological characterization of rhodopsin carriers
Both the DAB-labeled coated vesicles, either singly or in clusters (black arrows or curved arrows, Figure 6A–D), and DAB-labeled tubules (white arrows, Figure 6C, D) were abundant in the apical region of the rod inner segment. These tubular vesicle structures shared similar morphological features to those found in CC lumens, indicating that rhodopsin carriers seen in the CC were generated in the inner segments.
Finally, the ciliary PM and the apical inner segment PM of almost all transfected rods also had positive DAB reactivity (Figures 3B, C and 6A–G), consistent with the notion that rhodopsin is expressed on the PM of the CC.
Serial SEM examination of rod CC and membrane carriers
In almost all rods analyzed (n = 30), strings of vesicles were detected at the same site where the basal body lumen was localized (brackets in Figures 8, 9, and 10, Additional file 1: Movie 1) or exactly beneath that (white arrows in Figures 8, 9, and 10, black arrows in Additional file 1: Movie 1). These vesicles appeared to be part of a continuous flow of the vesicles in the CC inner shafts. Additionally, strings of vesicles were also seen near the base of the CP before they were channeled into the CC outer shafts (white arrowheads in Figures 8 and 10, white arrows in Figure 12, black arrows of Figure 9, and black arrowheads Additional file 1: Movie 1). In fact, CC vesicle strings were often connected to the vesicle strings localized in the upper inner segment (white arrows in Figures 10B and 12, black arrows in Additional file 1: Movie 1). Both en face (Figures 9, 10, and 13, white arrows in Additional file 1: Movie 1) and orthogonal (white arrows, Figure 13A) examinations showed some of these vesicles in the inner segments were immediately juxtaposed to a prominent organelle, which formed an extended tubular network spanning a long distance over the upper inner segment. These organelles were unlikely to be the trans-Golgi network, which is largely confined to the base of the inner segment . We were tempted to speculate that some of the CC vesicles were emanating from these as-yet unidentified membranous organelles.
The present study suggests that rhodopsin is translocated through the rod ciliary transition zone CC through multiple pathways, either on the PM or on membrane-bound carriers in the lumens. The membrane carriers, either large smooth vesicular tubules or small coated vesicles, are first synthesized in the inner segment, recruited at the base of the basal body, and then enter the inner shaft of the CC through the basal body lumen. Alternatively, they enter the CC outer shaft through the narrow cytoplasmic space between the AxMT/transition fibers and the ciliary PM.
Multiple types of rhodopsin transport carriers
While the previous cryo-EM report identified membrane carriers in the outer shaft, only large protein particles (~45–110 nm) were detected in the inner shaft of the CC . The conclusion was drawn based on the tomography analysis, which indicated that these large particles in the inner shaft had a different density profile compared to a typical phospholipid vesicle. On the other hand, several independent approaches in this paper all suggest the presence of CC membrane carriers in both the inner and outer shafts. First, our morphological characterization using high-resolution TEM reveals the presence of vesicles and tubules in the CC lumen. Second, our SEM analyses using a specific en bloc fixation/staining protocol reveals that the structures in CC lumens are membranous structures, not simple proteinous elements. The CC lumen vesicle density in the SEM images is evidently higher than that seen in the TEM images. We surmise it is due to the superiority of membrane preservation and membrane contrast due to the combinatory use of a stronger primary fixative and an en bloc post-fixation/staining protocol. Finally, we demonstrate the presence of integral membrane protein rhodopsin on both types of vesicular carriers in the CC lumen. The evidence includes (1) the immuno-EM reveals the expression of endogenous rhodopsin in the CC lumen and (2) DAB reaction products are specifically localized in the lumen side of the CC carriers in rods transfected with the topology-indicative HRP-rhodopsin reporter.
Our HRP-rhodopsin expression studies also reveal the expression of rhodopsin on the ciliary PM, in agreement with previous immuno-EM studies . Although the HRP-rhodopsin method has better accessibility than immuno-EM, we emphasize that no detergent was used in our experiments, in order to better preserve the membranes. So the HRP expressed inside the CC lumen is less accessible to the DAB substrates than that expressed on the outer surface. By the same token, we would like to reiterate that any given labeling protocol can only reveal a partial view of total protein expression. Despite using an anti-rhodopsin N-terminus Ab, Wolfrum and Schmit  showed only CC plasma membrane labeling, while our protocol can readily detect the rhodopsin inside the CC lumen using an anti-rhodopsin C-terminus Ab. The relatively low density of rhodopsin labeled in the CC lumen is expected, likely due to technical reasons, including Ab competition with the high concentration of rhodopsin in the OS and poor Ab penetration due to the dense extracellular matrix/large protein complex coating on the ciliary membranes. Note that for light microscopic examination, CC proteins have typically been labeled using unfixed or mildly fixed retinas [26,57,58], which are not suitable for ultrastructural analysis. Thus, we caution that the ratio of rhodopsin labeling on the ciliary PM vs. lumens (, current study) should not be directly interpreted as the relative amount of rhodopsin transported on different pathways.
Despite the relatively narrower space and the presence of Y-links, to some surprise, both the previous cryo-EM  and present studies reveal the existence of membrane carriers in the CC outer shaft. This finding, nonetheless, dovetails with the previous observation that excess vesicles accumulate in both inner and outer shafts of the rods that have a disc fusion problem .
Multiple trans-ciliary pathways in rod photoreceptors
While the biological meaning of having multiple types of rhodopsin carriers remains to be investigated, this finding suggests that various molecular motor systems might be involved in rhodopsin’s ciliary translocation. CC lumen-localized membrane carriers are within a close distance to the AxMT, making transport feasible through the use of the kinesin motor engaged on the AxMT (~25–30 nm; ). We speculate that the filamentous network linking the CC vesicles may be used to improve the processivity of motor-mediated transport activity and/or the efficiency of moving a large number of vesicles simultaneously. Filament-attached small vesicles have been seen in the developing mouse rod CC [10,60]. Obata et al.  showed that these filaments are myosin S1 fragment-decorated, short actin filaments; hence, actin-based motors (e.g., myosin VII [29,30]) may also be involved.
Coat formation is known to provide an effective means of concentrating membrane cargoes into patches; it is an evolutionarily ancient and conserved mechanism employed for selective sorting and preparing cargoes for transport from donor to recipient membranes . The coat composition of the CC vesicle is currently unknown, a piece of information important to further delineate the mechanism underlying the ciliary targeting and TZ entry of rhodopsin. In this vein, it is interesting to note that Arl6-mediated recruitment of Bardet-Biedl syndrome (BBS) protein can induce coated patches on liposomes, a process important for the primary ciliary entry . Mice with either the bbs2 or bbs4 gene deleted exhibited rhodopsin mislocalization [63-66], and bbs4 knockout mice also had aberrant vesicle accumulation at the CC base .
Regardless of the nature of the transport mechanism(s), the existence of multiple transport pathways for rhodopsin implies the possibility that partial, subtle, or even undetectable rhodopsin mislocalization may happen when a single translocation pathway is suppressed.
Rhodopsin’s disc incorporation and disc genesis
We imagine that the rhodopsin transported across the CC on the PM could diffuse into the OS PM, whereas that of transported on the membrane-bound carriers may directly fuse onto nascent discs. Supporting the latter notion, we found many rhodopsin-HRP carriers are in close proximity to the disc membranes, and the HRP labeling intensity of these carriers matches that of the disc membranes. Interestingly, some images show that rhodopsin is particularly concentrated on the edges of the membranous tubules in the basal OS axoneme (Figure 7A, B), indicating that the sorting continues even after these carriers have already entered the OS. We envision that the smaller vesicles with high rhodopsin density budded off from these tubules may be more fit for fusion to their immediately adjacent discs.
The above interpretation is in good agreement with the “vesicular targeting model” that we previously proposed for disc genesis . This model was initially proposed based on the observations that (1) the membrane fusion protein syntaxin 3 and membrane tethering protein Smad anchor for receptor activation are enriched at the most basal OS, the area in which the fusion activity is presumably the most robust, and (2) perturbing either molecule in rodent rods caused aberrant disc formation as well as vesicular accumulation inside the CC and OS base. According to this model, multiple nascent discs can be formed simultaneously at the basal OS via fusion of membrane carriers transported from the inner segment; repeated fusions allow the nascent discs to grow to the size of mature discs. This also contrasts to the “open disc/disc rim formation” model which suggests that a single disc is formed at a time. The present study using the topology-indicative rhodopsin reporter shows that both the mature discs and the nascent discs share the same membrane topology, and therefore, all should be enveloped by the OS PM in rat rods. This argues against the presence of evaginated or invaginated PM at the OS base.
Except for one TEM study that showed sparse vesicles in the lumen of the primary cilium of chondrocytes , no vesicles have been localized to simple (primary) cilia. Our findings here suggest that vertebrate rods might have evolved from other ciliated cells and developed a specialized means to move ciliary cargo in bulk due to the high demands posed by disc formation.
Transmission electron microscopy
Scanning electron microscopy
Serial block face scanning electron microscopy
Focused ion beam scanning electron microscopy
3,3′-Diaminobenzidine tetrahydrochloride hydrate
We thank Emily Beson for imaging collection and processing for SBF-SEM; William Rice and Edward T. Eng for operating and data collection of FIM-SEM (New York Structure Biology Center with the funding supported by NYSTAR, Research facilities Improvement Program Grant number C06 RR017528-01-CEM of National Center for Research Resources, NIH, NIH grant S10 RR029300). C-HS is supported by NIH grants (EY11307, EY016805) and Research to Prevent Blindness.
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