Multi-Modal Sample Preparation and Imaging Protocol for Nano-to-mesoscopic Mapping of Cellular Inhabitants in Diverse Tissue Compartments, Across Organ Systems

doi:10.21203/rs.3.pex-2244/v1

Method Article

Multi-Modal Sample Preparation and Imaging Protocol for Nano-to-mesoscopic Mapping of Cellular Inhabitants in Diverse Tissue Compartments, Across Organ Systems

https://doi.org/10.21203/rs.3.pex-2244/v1

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This protocol is designed to provide guidelines for preparation of samples that exceed typical sizes

used for standard microscopy protocols, expanding the methodologies and associated opportunities

for multimodal and correlative imaging to enable nano-to-mesoscopic mapping of cellular inhabitants

of diverse tissue compartments, across organ systems and length scales. We optimize and validate

specimen preparation protocols enabling multimodal and correlative imaging, LSM and scanning

electron microscopy (SEM), of large specimens of vertebrate animal origin. This protocol enables a

streamlined yet flexible approach to specimen preparation of large tissue samples (up to 10cm in

diameter). We take a novel approach to materials standard to EM, AFM, and confocal microscopy

specimen preparation, visualizing large fields of view without the need for heavy metal staining. The

protocol provides a platform for study of structure-function relationships across length scales, from

subcellular, cellular, tissue, to organ system. It is ideal for studies involving tissue specimens from

human or vertebrate animal models of human health and disease.

sample preparation

multimodal imaging

correlative imaging

cross length scales

cell to organ

meso scale

nano scale

Osteoarthritis (OA) is of the primary causes of disability in older adults and is characterized by chronic

pain and degeneration of synovial joint tissues ^1,2. Despite centuries of clinical and experimental study,

the precise mechanisms underpinning the onset of OA remain unknown ^2,3. This is reflected and

exacerbated by a dearth of FDA-approved biophysical, pharmaceutical (disease modifying

osteoarthritis drugs, DMOADs) and/or other means of disease prevention and intervention ².

Once regarded solely as a disease of the cartilage, current paradigms consider OA as a condition

involving all tissue compartments making up the synovial joint organ, including synovium, cartilage,

bone, subchondral bone, periarticular musculature, meniscus, ligament, and other connective joint

tissues ⁴. Rather than considering OA as a product of the aging process and mechanical ‘wear and

tear’, the onset and progression of OA is now seen as a consequence of intertwined biomechanical

and biochemical processes, leading to the eventual failure of the joint ⁵. Yet research often focuses on

singular facets of the disease, e.g. cellular interactions, disease processes, or risk factors associated

with OA. An understanding the structure-function relationships within the complex biosystem of the

joint, spanning multiple lengths scales from the molecular, cellular, tissue, organ to organ system. is

imperative to decipher the pathogenesis of OA and to develop therapeutic targets preventing its onset

and/or progression ^6-8.

Confocal laser scanning microscopy (LSM) has proven to be a crucial tool for unravelling cell

physiology, and enabling visualization of highly dynamic biological processes ⁹. Similarly, transmission

electron microscopy (TEM), provides unique advantages for resolving the complexity of cells and their

subcellular structures and thus serves as a seminal technology for cell biology ¹⁰. However, LSM and

TEM imaging modalities rely on often laborious samples preparation techniques and typically limit

specimen size to cells or small animals (e.g. murine specimens) or subsections of tissue that do not

exceed several millimeters on edge. The development of automated tiling and stitching software for

LSM has enabled imaging of large fields of view at high resolution of specimens up to 67 x 120mm.

The advent of multibeam scanning electron microscopy (multiSEM) enables the imaging of large

specimens, up to 100 mm in diameter, at nm resolution, far exceeding the capabilities of traditional

single-beam SEM. Interdisciplinary research and the trans-dimensional linking of biological structure-

function relationships are propelled by the development of seamless, multi-scale and multi-modal

imaging platforms ^11–22. With the emergence of these new imaging and image processing technologies,

parallel advances in specimen preparation techniques are needed.

Here we optimize and validate specimen preparation protocols enabling multimodal and correlative

imaging, LSM and scanning electron microscopy (SEM), of large specimens of vertebrate animal

origin. These protocols allow multiscale and multimodal visualization of structure-function relationships

of large tissues for the elucidation of joint physiology in health and disease states.

Materials and Methods

Sample size guidelines

The protocol is designed to provide guidelines for preparation of samples that exceed typical sizes

used for standard microscopy protocols, expanding the methodologies and associated opportunities

for multimodal and correlative imaging to enable nano-to-mesoscopic mapping of cellular inhabitants

of diverse tissue compartments, across organ systems and length scales. In previous studies, human

femoral neck, and head specimens, obtained from total hip arthroplasty patients, were sectioned into 1

mm thick sections along the plane of interest by the Cleveland Clinic Pathology Department per IRB

protocol (#12-335) guidelines and the resulting large scale tissue blocks were imaged, from nm to

meso-length scales using multibeam scanning electron microscopy ^{11,16-20,22,23}. The current study further

develops the protocol for multimodal and correlative imaging across length scales using Dunkin

Hartley guinea pigs as a model system for age-related tissue degeneration, e.g. associated with

osteoarthritis.

Experiment and Specimen preparation (see Figure 1, first series of blue arrow shapes)

Animal experiments were approved by Case Western Reserve University IACUC (Protocol #2012-

0100) and were performed in accordance with the ARRIVE guidelines and regulations. In all

previously published experiments from our laboratory using the guinea pig model, a bolus of

fluorescent tracer was administered via the heart ^6-8. In the current study ⁷, a bolus of 70 kDa fluorescent

tracer was administered intracardially according to our previously published protocols; two

experimental groups either included one of two cytokines, TGF-alpha or TNF-beta, mixed in the bolus of

fluorescent tracer. After five minutes' circulation, animals were euthanized and hindlimbs were

immediately resected (Figure 1, Sample Acquisition, Sectioning, and Fixation).

Tissues were immediately either cryopreserved in OCT using liquid nitrogen for Episcopic

Cryoimaging (Figure 1) ^6-8 or chemically fixed⁷ in 2.5% glutaraldehyde and 4% paraformaldehyde in 0.2

M cacodylate buffer for five days. Thereafter, samples were washed with cacodylate buffer daily for

another five days. Samples were then dehydrated in a series of ethanols (30%, 50%, 75%, 100% -

Figure 1, Dehydration) for three days each with daily solution changes. After dehydration, samples

were embedded in polymethylmethacrylate (PMMA, Figure 1 - Resin Embedding) resin.

Surface Preparation

In the present study, samples were micro-CNC milled to obtain a mirror-like finish in areas to be

imaged. CNC-milled samples were fixed onto a CNC-lathe and milled with a carbide insert, followed

by gentle buffing with an acrylic polish. Then, the samples were gently cleaned with ultra-pure H2O

then absolute ethanol with a lint free wipe and were similarly placed under a stream of inert gas.

For laboratories that do not have access to micro-CNC milling, machine polishing is an alternative

method that we have tested in other studies using a semi-automatic grinder and polisher (Struers

LaboPol-5). For machine polishing, specimens are placed, with even pressure, surface-side down on

a 500-grit polishing disc at 200 rpm and then cleaned ultrasonically in a beaker of deionized water.

This process is repeated with a 1200-grit polishing disc, MD-Dac polishing pad with 3 micron diamond

suspension, and then MD-Nap polishing pad with 1 micron diamond suspension. Following the polishing

steps, specimens are gently cleaned with absolute ethanol and a lint-free wipe, then placed under a

steady stream of inert gas, in this case nitrogen, to ensure they are dry and free of any particulates.

After surface preparation, specimens can be directly imaged using confocal imaging and multiphoton

imaging modalities (see below). If multimodal imaging is planned, e.g. confocal and electron

microscopy imaging, is planned, then it is best to first carry out contrast enhancement, e.g. with

chemical etching or iodine staining, prior to confocal imaging and electron microscopy (Figure 1).

Contrast enhancement and viability assay with iodine

To enhance contrast and resolve cellular detail using electron microscopy (EM), specimens

underwent chemical etching and/or iodine staining.

We used chemical etching to reveal osteocyte networks in calcified tissues, adapting protocols

that had been previously implemented for atomic force microscopy of osteocyte lacunocanalicular

networks in bone ²⁴. Specimens were mounted onto a specimen holder using silver paint, carbon,

or copper tape, then gently cleaned with absolute ethanol and a lint free wipe. The sample was

then placed under a stream of inert gas to ensure it was dry and free for dust. A small volume of

0.02 M hydrochloric acid (HCl) was pipetted onto the specimen surface, then a cotton swab was

used to gently expose the sample to the solution for 60 seconds at time, the sample was then

flushed with ultra-pure H2O to clear the surface of any debris and residual solution. Following this,

a similarly small volume of 2.5% sodium hypochlorite (NaClO) was applied to the specimen in the

same manner as the previous step, including flushing with ultra-pure H2O. Specimens were then

placed under a steady stream of inert gas to ensure that they were completely dry and free of

dust then carbon coated (20nm thickness) in preparation for electron microscopy (EM).

Boyde et al. described the use of iodine staining as an assay for cell viability and we adapted

Boyde's protocol for the current study ^25,26. Specimens undergoing iodine staining were cleaned

with absolute ethanol and a lint free wipe then placed under a stream of inert gas prior to carbon

coating (20 nm thickness). Lugol solution (aqueous potassium iodide and iodine) was then placed

on the sample surface for 5 minutes to impart an iodine stain. The excess Lugol solution was

then flushing with ultra-pure H2O and the specimen was placed under a steady stream of inert

gas.

Multimodal imaging

We imaged specimens first using confocal laser scanning microscopy (LSM); for the current study we

used a Zeiss LSM 880, an inverted laser scanning confocal microscope equipped with an Argon laser

for 488 nm excitation and Helium-Neon (HeNe) laser for 594 excitation, 10Å~ 0.8 Plan-Apochromat

dry objective, 20Å~ 0.8 Plan-Apochromat dry objective and 63Å~ 1.4 Plan-Apochromat oil objective to

observe tissue autofluorescence and the Texas-red fluorophore.

After completion of confocal imaging, in the current study we then carried out backscattered electron

scanning electron microscopy (BSE-SEM) or secondary electron scanning electron microscopy (SEM)

using an FEI NanoSEM 450. In previous studies ^16-20,22,23, specimens underwent widefield light

microscopy, using a Zeiss Axio Imager Vario, followed by rapid throughput, secondary electron

scanning electron microscopy, using a Zeiss multiSEM platform, with up to 91 parallel electron beams.

Results

Fixation and Resin Embedding

Use of PMMA resin embedding, together with the exclusion of heavy metal (OsO₄) staining, enabled the preparation of optically clear specimens conducive to light microscopy. Upon imaging, separation artifacts were observed as empty spaces along tissue compartment boundaries where attached tissues had pulled away from each other, due to shrinkage during chemical fixation ²⁷. The artifacts were observed at the proximate to the boundary between bone and bone marrow, along bounding fasciae and between muscle and bone; no separation was observed at the osteochondral interface (Figure 2).

Surface Preparation

While a mirror-like finish is achievable using semi-automatic polishing, some specimens develop micro-cracks on the sample surface. These microcracks often appear at tissue boundaries, causing loss of data, and interfering with both light microscopy and EM imaging quality. Further, using semi- automatic polishing, it is challenging to achieve planarity required for imaging large regions, e.g. for multibeam scanning electron microscopy. CNC-milling, followed by manual buffing, enables both high-precision removal of material as well surface planarity without the development of micro-cracks which interfere with image quality.

Chemical Etching

Chemical etching is necessary for resolving cellular detail in sample bloscks using SEM. The combined action of dilute sodium hypochlorite and hydrochloric acid enables controlled erosion of the organic and inorganic apatite matrix components, respectively, exposing osteocytes and osteocyte processes embedded in the extracellular matrix (ECM, Figure 3).

Iodine staining

While carbon coating alone provided sufficient contrast to visualize bone and reveal the osteocyte lacuno-canalicular network (LCM) on the sample surface, other tissues remained dark with no discernable features (Figure 4). Lugol solution imparted an iodine stain to musculoskeletal tissues with little to no mineralization, enabling visualization of previously undiscernible musculoskeletal tissues including articular cartilage, calcified cartilage, growth plate cartilage, muscle, periosteum, and bone marrow, in addition to bone. Subcutaneous adipose tissue, arteries and veins were also stained. Osteocytes and the LCM appeared dark against bone with nuclei exhibiting slightly elevated levels of stain in comparison to the cytoplasm. Chondrocytes were stained in a similar manner; however articular cartilage was stained less intensely than bone. Some subcutaneous adipocyte nuclei were observable, appearing brighter than the cytoplasmic rim; no stain was imparted on stored lipids. Marrow cells appeared slightly brighter than the marrow space, bone marrow adipocytes appeared darker than surrounding marrow (Figure 5).

Multi-Modal Imaging

The initial generation of an overview image of the sample enabled the isolation of regions of interest for further imaging (Figure 3D; see Figures in related articles ^7,8); a low-resolution image is sufficient for this purpose. This also allows for the cross-identification of landmarks for further multi-modal imaging and correlation. The addition of extrinsic fiduciaries to the sample such as a small score to the edge of the specimen aided with navigating large specimens, especially on subsequent imaging modalities. It was important to take note of and record the orientation of the specimen as it is placed onto the microscope stage as well as any changes in rotation in the resultant image. We observed that the resultant laser scanning microscopy and SEM images were mirrored horizontally and rotated 90° clockwise, respectively.

Discussion

Advances enabling high throughput imaging, such as the multiSEM platform and tiling and stitching software, have made it possible to perform SEM and LSM of specimens up to 10 cm in diameter at nanometer and micron length scales ^{11,16-20,22,23}. In this present study we expanded upon specimen preparation techniques ordinarily used for TEM and SEM, enabling visualization of tissues from the nano-to-mesoscopic length scales. We adapted chemical etching methods, developed for atomic force microscopy, and previously described iodine staining to facilitate multi-modal and correlative imaging ^24-26. These methods were applied previously to large areas of human tissue (sections of femoral head and neck) and the current protocol highlights their use in tissues of medium-sized vertebrates (guinea pig knee joints including surrounding musculature), demonstrates their feasibility for integration with multi-modal imaging workflows, enabling correlative studies of structure and function spanning multiple length scales.

While human tissues present a gold standard for study of human health and mechanisms underpinning disease, intrinsic limitations of human tissue relate to both contexts for ethical implementation and associated lack of experimental controls; while the tissues used in previous studies of the human hip would normally be discarded in the typical course of joint replacement, resection of healthy tissues to serve as a baseline control would be neither ethical nor tenable. As such, animal models offer an alternate pathway for deciphering pathophysiologic mechanisms underpinning disease states and associated loss in tissue/cell structure and function in humans. Pairing animal models with human tissue studies provides a powerful means to understand the human condition in parallel vertebrate animal models, and establishment of methods that apply for both human and animal tissues further amplifies that power.

The Dunkin-Hartley guinea pig exhibits age-related spontaneous osteoarthritis which mirrors the condition in humans ²⁸. Previous studies implementing the Dunkin-Hartley model have demonstrated the functional barrier property of molecular sieving in musculoskeletal tissues and passive transport of large molecules modulated by age and inflammatory state ^6-8. Through cross length scale imaging using multiple imaging modalities – such modalities include episcopic meso to cell-scale imaging of the entire lower limb, high resolution confocal imaging of the same mesoscopic regions in subcellular resolution, and electron microscopy – unprecedented mechanistic relationships were drawn between the cardiovascular and musculoskeletal systems in association with osteoarthritis. In particular, cardiovascular transport from the heart to the musculoskeletal system, could be visualized as a function of molecular size and exposure to immunomodulatory cytokines, using fluorescent probes that cannot be used in humans. Together, these studies provide a new basis of understanding for structure-function relationship between tissues and the cells that inhabit them, across organ systems.

Whilst study of single types of molecules, cells, tissues, organs, and organ systems has yielded a plethora of scientific knowledge, new discoveries are predicted to occur at interfaces between organ systems/tissues and through interactions amongst living entities, e.g. cells ¹¹. Hence, here we expand upon this specimen preparation protocol to enable study of the interplay between and across length scales, organ systems (cardiovascular to musculoskeletal to immune systems), organs, tissues and cells ^6-8,23. The protocol enables unprecedented epidemiological approaches to understand organismal health, be it human or guinea pig, in context of the health of cell populations inhabiting tissues, organs and organ systems.

Heavy metal post-fixation staining is typically required to provide sufficient contrast to tissue specimens and to resolve cellular detail for EM ^29,30. The use of heavy metals for contrast enhancement, e.g. typically uranyl acetate, lead citrate and osmium tetroxide, is often a labor-intensive process and requires exposure to extremely hazardous chemicals. Osmium tetroxide imparts an exogenous stain on the entire specimen, reducing optical clarity and hindering light microscopy. Uranyl acetate and lead citrate introduce instability into the specimen, with the risk of precipitation artifact formation over time ³¹.

The substitution of heavy metal contrast agents for post-hoc chemical etching and iodine staining, in addition to carbon sputtering, provides sufficient contrast and eliminates the use of hazardous chemicals and their associated risks while maintaining sample stability. Chemical fixation of cells and tissues has been shown previously to result in anisotropic shrinkage of cells ²⁷. In addition, tissue dehydration may result in a 5–70% reduction of tissue volume, resulting in separation artefacts ³². Dehydration involves organic solvents and is vital to the resin embedding process, as PMMA is not miscible in water. Furthermore, separation artifacts have been observed between bone and bone‐anchored metal implants, suggesting that such artifacts arise at the interface of materials and/or tissues of significantly dissimilar physical properties ³³. This is consistent with our observation of separation artifacts at the tissue boundaries. Separation artifacts caused by dehydration can be reduced by embedding specimens in more water-miscible resins and through the use of cryo-embedding methods such as high-pressure freezing and freeze substitution ^34–38. However, it should be noted that while cryo-preservation is a well-established technique for cells and small tissue volumes, cryo-preservation of bulk tissues remains difficult; even with the use of cryo-protective agents, the formation of extracellular ice crystals can disrupt tissue structures ³⁸.

The surface flatness of specimens is vital to EM; defects such as micro-cracks generate edge charging artefacts, causing losses in data, resolution and may result in specimen damage ³⁹. CNC-milling proved to be the superior method for achieving planarity and a mirror-like finish on the specimen surface. Semi-automated surface preparation proved to be a laborious and time-intensive process. In addition to this, we observed the development of micro-cracks in the sample surface, possibly due to release of residual stresses generated during the resin embedding process. Unlike smaller, conventionally sized, samples, the specimens could not be tilted within the microscopes to level the specimen surface for imaging. While more cautious and slower manual polishing could potentially minimize microcracks, manual surface preparation techniques are not as conducive to achieve the levels of parallelism between the top and bottom surfaces of the sample required for imaging.

While the flexibility of this method enables specimens to be imaged using multiple platforms, this same feature makes it not ideal for integrated correlative light and electron microscopy system as the specimen must be removed from the microscope chamber to undergo additional contrast enhancement steps after light microscopy to make it conducive to EM. This protocol enables a streamlined yet flexible approach to specimen preparation of large tissue samples (up to 10cm in diameter). We take a novel approach to materials standard to EM, AFM, and confocal microscopy specimen preparation, visualizing large fields of view without the need for heavy metal staining. The protocol provides a platform for study of structure-function relationships across length scales, from subcellular, cellular, tissue, to organ system. It is ideal for studies involving tissue specimens from human or vertebrate animal models of human health and disease.

1. World Health Organisation. Disease incidence, prevalence and disability. The Global Burden of

Disease: 2004 Update (2008).

2. Ngo, L., & Knothe Tate, M.L. Osteoarthritis: New strategies for transport and drug delivery, ACS

Biomaterials Science & Engineering, 6: 6009 - 6020 (2020).

https://doi.org/10.1021/acsbiomaterials.0c01081.

3. Dequeker, J. & Luyten, F.P. The history of osteoarthritis-osteoarthrosis. Ann. Rheum. Dis. 67, 5–10

(2008).

4. Loeser, R.F., Goldring, S.R., Scanzello, C.R., & Goldring, M.B. Osteoarthritis: A disease of the joint

as an organ. Arthritis Rheum. 64, 1697–1707 (2012).

5. Aigner, T. & Schmitz, N. Pathogenesis and pathology of osteoarthritis. in Rheumatology (ed.

Hochberg M, Silman A, Smolen J, Weinblatt M, W. M.) 1741–1759 (Mosby Elsevier, 2011).

6. Ngo, L., Knothe, L., & Tate M.L.K., Knee Joint Tissues Effectively Separate Mixed Sized Molecules

Delivered in a Single Bolus to the Heart, Nature Scientific Reports, 8: 10254(1-6) (2018).

doi:10.1038/s41598-018-28228-w

7. Ngo, L, & Knothe Tate, M.L. A spike in circulating inflammatory cytokines alters barrier function

between vascular and musculoskeletal tissues, Nature Scientific Reports, doi:10.1038/s41598-023-30322-7 (2023).

8. Ngo, L, & Knothe Tate M.L. Inflammatory cytokines modulate transport across length scales. from

heart to joint, tissues to cells, Orthopaedic Research Society Meeting, 46: 0604 (2021).

9. Paddock, S.W. & Eliceiri, K.W. Laser Scanning Confocal Microscopy: History, Applications, and

Related Optical Sectioning Techniques. in 9–47 (2014). doi:10.1007/978-1-60761-847-8_2.

10. Harris, J.R. Transmission electron microscopy in molecular structural biology: A historical survey.

Arch. Biochem. Biophys. 581, 3–18 (2015).

11. Knothe Tate, M.L. Navigation of bee brains to human hips - microscopy and the modern

Magellans. in A new age in scanning electron microscopy: Applications in the life sciences 19–23

(Science/AAAS, 2017).

12. Sporns, O., Tononi, G., & Kötter, R. The human connectome: A structural description of the human

brain. PLoS Computational Biology vol. 1 0245–0251 (2005).

13. National Institutes of Health. NIH to build a detailed map of cells within the human body. (2018).

14. Lichtman, J.W. & Sanes, J.R. Ome sweet ome: what can the genome tell us about the

connectome? Curr. Opin. Neurobiol. 18, 346–353 (2008).

15. Knothe Tate, M.L., Detamore, M., Capadona, J.R., Woolley, A., & Knothe, U.R. Engineering and

commercialization of human-device interfaces, from bone to brain. Biomaterials 95, 35–46 (2016).

16. Eberle, A.L., Mikula, S., Schalek, R., Lichtman, J., Tate, M.L.K., Zeidler, D., & Knothe Tate, M.L.

High-resolution, high-throughput imaging with a multibeam scanning electron microscope. J. Microsc.

259, 114–20 (2015).

17. Knothe Tate, M.L., Zeidler, D., Pereira, A.F., Hageman, D., Garbowski, T., Mishra, S., Gardner, L.,

& Knothe, U.R. Organ-to-Cell-Scale Health Assessment Using Geographical Information System

Approaches with Multibeam Scanning Electron Microscopy. Adv. Healthc. Mater. 5, 1581–1587 (2016).

18. Hageman, D., Pereira, A.F., Zeidler, D., Knothe, U.R., Gardner, L., & Knothe Tate, M.L. Cellular

Epidemiology of Human Disease using Biogeographic and Google Maps Approaches - Towards

Definition of Cell Network Indices for Rapid Diagnostics. Ann. Biomed. Eng. (2016).

19. Tate, M.L.K. & Fath, T. The Only Constant Is Change: Next Generation Materials and Medical

Device Design for Physical and Mental Health. Adv. Healthc. Mater. 5, 1840–1843 (2016).

20. Pereira, A.F., Hageman, D.J., Garbowski, T., Riedesel, C., Knothe, U.R., Zeidler, D., & Knothe

Tate, M.L. Creating High-Resolution Multiscale Maps of Human Tissue Using Multi-beam SEM. PLoS

Comput. Biol. 12, e1005217 (2016).

21. National Institutes of Health. The Human Connectome Project.

https://www.humanconnectome.org/.

22. Knothe Tate, M.L., Srikantha, A., Wojek, C., & Zeidler, D. Connectomics of Brain to Bone - Probing

Physical Renderings of Cellular Experience, Frontiers in Physiology | Integrative Physiology, Special

issue on the relationship between neural circuitry and biomechanical action, 12:647603 (2021), doi:

10.3389/fphys.2021.647603

23. Ngo, L., Nathanson, A., Garbowski, T., Knothe, U., Zeidler, D., & Knothe Tate, M.L. Electron

microscopy sample preparation protocol enabling nano-to-mesoscopic mapping of cellular

connectomes and their habitats in human tissues and organs, Bio-protocol, 9(14): e3298(1-18),

(2019). doi: 10.1101/533448

24. Knapp, H.F., Reilly, G.C., Stemmer, A., Niederer, P., & Knothe Tate, M.L. Development of

preparation methods for and insights obtained from atomic force microscopy of fluid spaces in cortical

bone. Scanning 24, 25–33 (2002).

25. Boyde, A. Staining plastic blocks with triiodide to image cells and soft tissues in backscattered

electron SEM of skeletal and dental tissues. Eur. Cells Mater. 24, 154–161 (2012).

26. Boyde, A., Mccorkell, F.A., Taylor, G.K., Bomphrey, R.J., & Doube, M. Iodine vapor staining for

atomic number contrast in backscattered electron and X-ray imaging. Microsc. Res. Tech. 77,

1044–1051 (2014).

27. Zimmerman, J., & Knothe Tate, M.L. Structure - function relationships in the stem cell's mechanical

world A: Seeding protocols as a means to control shape and fate in live cells, Molecular and Cellular

Biomechanics, 8(4): 275-296 (2011). eid: 2-s2.0-84555217837

28. Jimenez, P.A., Glasson, S.S., Trubetskoy, O. V, & Haimes, H.B. Spontaneous osteoarthritis in

Dunkin Hartley guinea pigs: histologic, radiologic, and biochemical changes. Lab. Anim. Sci. 47,

598–601 (1997).

29. Watson, M.L. Staining of Tissue Sections for Electron Microscopy with Heavy Metals. J. Biophys.

Biochem. Cytol. 4, 475–478 (1958).

30. Wigglesworth, V.B. The use of osmium in the fixation and staining of tissues. Proc. R. Soc.

London. Ser. B - Biol. Sci. 147, 185–199 (1957).

31. Pandithage, R. & Leica Microsystems GmbH. EM Sample Preparation Contrasting. (2013).

32. Mollenhauer, H.H. Artifacts caused by dehydration and epoxy embedding in transmission electron

microscopy. Microsc. Res. Tech. 26, 496–512 (1993).

33. Palmquist, A., Lindberg, F., Emanuelsson, L., Brånemark, R., Engqvist, H., & Thomsen, P.

Morphological studies on machined implants of commercially pure titanium and titanium alloy

(Ti6Al4V) in the rabbit. J. Biomed. Mater. Res. Part B Appl. Biomater. 91B, 309–319 (2009).

34. Peddie, C.J., Liv, N., Hoogenboom, J.P., & Collinson, L.M. Integrated Light and Scanning Electron

Microscopy of GFP-Expressing Cells. in Methods in Cell Biology 363–389 (2014). doi:10.1016/B978-0-12-801075-4.00017-3.

35. Johnson, E., Seiradake, E., Jones, E.Y., Davis, I., Grünewald, K., & Kaufmann, R. Correlative in-

resin super-resolution and electron microscopy using standard fluorescent proteins. Sci. Rep. 5, 9583

(2015).

36. Watanabe, S., Punge, A., Hollopeter, G., Willig, K.I., Hobson, R.J., Davis, M.W., Hell, S.W., &

Jorgensen, E.M. Protein localization in electron micrographs using fluorescence nanoscopy. Nat.

Methods 8, 80–84 (2011).

37. Bullen, A., Taylor, R.R., Kachar, B., Moores, C., Fleck, R.A., & Forge, A. Inner ear tissue

preservation by rapid freezing: Improving fixation by high-pressure freezing and hybrid methods. Hear.

Res. 315, 49–60 (2014).

38. Pegg, D.E. Principles of Cryopreservation. in (eds. Wolkers, W. F. & Oldenhof, H.) vol. 1257 3–19

(Springer New York, 2015).

39. Kjellsen, K.., Monsøy, A., Isachsen, K., & Detwiler, R.. Preparation of flat-polished specimens for

SEM-backscattered electron imaging and X-ray microanalysis—importance of epoxy impregnation.

Cem. Concr. Res. 33, 611–616 (2003).

Author contributions

The specimens were obtained from M.K.T from previous guinea pig and human studies under respective institutional ethics approvals. Chemical etching enhancement techniques were implemented and multi-modal imaged and analyzed by L.N. Testing and implementation of iodine contrast enhancement of guinea pig specimens; high resolution confocal imaging, tiled image mapping and correlative imaging was conducted by L.N. Writing of the protocol was carried out by L.N. and editing of the protocol was carried out by L.N. and M.K.T.

Acknowledgements

We would like to express our special appreciation to Professor Alan Boyde of the Institute of Dentistry, Queen Mary University of London, for fruitful discussions and for his advice regarding shortfalls of morphology-based etching and associated advantages of implementing the iodine staining portion of the protocol for determination of cell presence and viability. We would also like to acknowledge the clinical and surgical expertise of Dr. Ulf Knothe and Dr. Thomas Bauer for specimen acquisition under the IRB protocol (Cleveland Clinic Foundation), scientific and technical assistance, as well as the the Core Research Services of the Lerner Research Institute (Cleveland Clinic Foundation, Dr. Judith Drazba, Ms. Mei Lin). Mr. Anthony Shawan of the Cleveland Clinic carried out the precision CNC milling and polishing. Multibeam-SEM imaging was carried out in the demonstration labs of Zeiss Microscopy GmbH in Oberkochen, Germany, with in kind support from Dr. Dirk Zeidler and his team. Confocal imaging, TPIM and SHIM, as well as single beam electron microscopy were carried out at University of New South Wales' Microscopy Australia at the Electron Microscope Unit (EMU) and the Biomedical Imaging Facility (BMIF) within the Mark Wainwright Analytical Centre (MWAC).

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