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Crystal ribcage: a platform for probing real-time lung function at cellular resolution

Nature Methods volume 20, pages 1790–1801 (2023)Cite this article

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Abstract

Understanding the dynamic pathogenesis and treatment response in pulmonary diseases requires probing the lung at cellular resolution in real time. Despite advances in intravital imaging, optical imaging of the lung during active respiration and circulation has remained challenging. Here, we introduce the crystal ribcage: a transparent ribcage that allows multiscale optical imaging of the functioning lung from whole-organ to single-cell level. It enables the modulation of lung biophysics and immunity through intravascular, intrapulmonary, intraparenchymal and optogenetic interventions, and it preserves the three-dimensional architecture, air–liquid interface, cellular diversity and respiratory–circulatory functions of the lung. Utilizing these capabilities on murine models of pulmonary pathologies we probed remodeling of respiratory–circulatory functions at the single-alveolus and capillary levels during disease progression. The crystal ribcage and its broad applications presented here will facilitate further studies of nearly any pulmonary disease as well as lead to the identification of new targets for treatment strategies.

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Fig. 1: Development, capabilities and application of the crystal ribcage in pulmonary research.
Fig. 2: Multiscale structure–function changes in lung cancer models.
Fig. 3: Impairment of alveolar respiratory and circulatory functions in pneumonia.
Fig. 4: Neutrophil migration is reversibly mechano-responsive to vascular pressure.
Fig. 5: Respiration–circulation coupling in healthy lungs and its disruption by acute injury.

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Data availability

A demonstration dataset has been deposited in a Zenodo repository82 and is available from https://doi.org/10.5281/zenodo.7939072. Additional microscopy data are available on request from the corresponding author. Source data are provided with this paper.

Code availability

The source code and algorithms used in this study have been deposited in a Zenodo repository82 and are available from https://doi.org/10.5281/zenodo.7939072 licensed under the CC BY 4.0 for use in research.

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Acknowledgements

We thank staff at the Neurophotonics Center at Boston University for their generous support and access to their facility. A. Coats from the Hoffman laboratory at the University of Iowa provided the µCT image datasets used to fabricate the crystal ribcage. The research reported in this publication was supported by the Boston University Micro and Nano Imaging Facility and the Office of the Director, National Institutes of Health, under award no. S10OD024993. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Any opinion, findings and conclusions or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the National Science Foundation. H.T.N. discloses support for the research described in this study from the National Institutes of Health R21EB031332 and DP2HL168562, a Beckman Young Investigator Award, an NSF CAREER Award, the Johnson & Johnson Lung Cancer Initiative, Boston University Center for Multiscale and Translational Mechanobiology and a Dean’s Catalyst Award and the American Cancer Society Institutional Fund at Boston University. L.S. acknowledges support from the National Heart, Lung and Blood Institute grant 1F30HL168952-01. This material is also based upon work supported by the National Science Foundation and G.N.G. acknowledges support from the NSF Graduate Research Fellowship, grant no. 2234657. The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.

Author information

Author notes

  1. These authors contributed equally: Rohin Banerji, Gabrielle N. Grifno.

Authors and Affiliations

  1. Department of Biomedical Engineering, Boston University, Boston, MA, USA

    Rohin Banerji, Gabrielle N. Grifno, Linzheng Shi, Dylan Smolen, Rob LeBourdais, Johnathan Muhvich, Cate Eberman, Kathryn Regan, Siyi Zheng, Sue Zhang, John Jiang, Béla Suki & Hadi T. Nia

  2. Pulmonary Center, Boston University Chobanian & Avedisian School of Medicine, Boston, MA, USA

    Bradley E. Hiller, Jisu Lee, Ahmed A. Raslan, Riley Pihl, Katrina Traber, Giovanni Ligresti, Joseph P. Mizgerd & Hadi T. Nia

  3. Department of Zoology, Faculty of Science, Assiut University, Assiut, Egypt

    Ahmed A. Raslan

  4. Section of Computational Biomedicine, Boston University Chobanian & Avedisian School of Medicine, Boston, MA, USA

    Julia C. Breda & Sarah Mazzilli

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Contributions

R.B., G.N.G. and H.T.N. conceived the project and wrote the manuscript; R.B. developed and validated the crystal ribcages and external devices suite, established transgenic mouse colonies and analyzed alveolar function, strain maps and collagen waviness; G.N.G. conducted the lung extraction and imaging, generated experimental mice, cultured cancer cells and analyzed circulatory function data; L.S. analyzed perfusion, vascular transport and neutrophil mechano-responsiveness data; D.S. analyzed alveolar and capillary function data and maintained the animal colony; R.L. developed the registration and strain analysis algorithm; J.M. developed custom hardware for crystal ribcage and analyzed alveolar function data; C.E. developed the deformable crystal ribcage; B.E.H. generated the lobar pneumonia mouse model; J.L. and A.A.R. generated the fibrosis model; A.A.R. performed the histology and staining for fibrotic lungs; J.C.B. generated primary cancer mice, K.R. analyzed alveolar function data and cultured cancer cells; S. Zheng cultured cancer cells; S. Zhang fabricated fluorescent polyacrylamide beads; J.J. assisted in surgical techniques for in situ lung imaging; R.P. generated the LPS induction model; K.T. provided the transgenic MPR8-Cre+ mouse; S.M., G.L., J.P.M. and B.S. contributed to discussion on crucial aspects of the project; H.T.N. supervised the project and provided guidance on experimental design, data collection, data interpretation and writing of the manuscript.

Corresponding author

Correspondence to Hadi T. Nia.

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Competing interests

H.T.N. has received research funding from Johnson & Johnson Lung Cancer Initiative at Boston University. The funding has supported the development of the lung cancer models described in the manuscript. All other authors report no competing interests.

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Nature Methods thanks Joan Nichols and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling Editor: Madhura Mukhopadhyay, in collaboration with the Nature Methods team.

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Extended data

Extended Data Fig. 1 Mouse strain-specific µCT informed 3D molds scaled by age and fabrication of crystal ribcages.

(a) Recreating mouse ribcage geometry from µCT data75,76,77, here shown for C57/B6 mice. Top: Sagittal view of mouse chest with fully inflated lung (dark region), distribution of axial control points to recapitulate ribcage cross-section and comparison of reconstructed geometry overlaid on sagittal and coronal µCT slices. Lung volume variation78 with age was normalized against the mean volume at 3 months to construct a linear fit for scaling 3D lung inserts by age. (b–e) 3D printed models were used to make both PDMS and polystyrene crystal ribcages in two independent processes74.

Extended Data Fig. 2 Native mouse ribcage does not show significant variation in diameter across its anatomy and alveolar function remains unchanged between crystal ribcage and mouse ribcage.

Previous µCT data75,76,77 used to define the mouse chest cavity was also used to quantify the expansion of the native mouse ribcage as a function of pressure and anatomical location during in situ ventilation. There was no significant difference between the ribcage diameter at tracheal pressures ranging from 10 to 40 cmH2O in both the frontal (a) and sagittal (b) planes from the apex to the base of the ribcage. The increase in volume with increasing pressure was accommodated by the increasing length of the lung (movement of diaphragm and abdomen) and not the width of the ribcage. This confirms that there was little contribution of the changing ribcage diameter to lung volume change during normal physiological conditions in situ. The scatter of the data over n = 3 mice is presented along with the mean ± SEM traces. (c) A mouse ribcage was thinned down to make a window above the pleural sheath to image the functional lung using OCT, the same lung was then measured in the crystal ribcage for the same changes in pressure. There was no significant difference between alveoli function (measured as changing diameter) across the pressure range for m=5 ROI over n = 1 mouse. The scatter of the data is presented along with the mean ± SEM traces. Comparison between groups performed with two-tailed Student’s t-test for significance < 0.05.

Source Data

Extended Data Fig. 3 Experimental setup and 3D render of the microscopy arm used to image different orientations of the crystal ribcage with upright and inverted microscope configurations.

We successfully used the stage to image (a) with an inverted Olympus Fluoview 3000 laser-scanning confocal microscope and (b) with upright microscopes, such as the ThorLabs optical coherence tomography probe and Bruker multi-photon microscope. The crystal ribcage is microscope agnostic and compatible with both air and water immersion objective in both configurations.

Extended Data Fig. 4 Dynamic ventilation of mouse lung shows alveoli remain at a larger diameter for longer duration at higher ventilation rate.

Comparing the same region of the lung ventilated at (a) 60 breaths/minute with (b) 120 breaths/minute by (c) the mean ± SEM alveolar diameter from m=4 ROI over n = 2 mice with a normalized ventilation cycle. Blue traces in the inset images show the segmentation boundary drawn to quantify single alveoli. (d) The width of the normalized ventilation cycle from m=4 ROI over n = 2 mice at each respiratory rate are statistically larger at the higher respiratory rate. Boxplots present median with 25th and 75th percentiles, whiskers are the maximum and minimum data points not considered outliers. Comparison between groups performed with two-tailed Student’s t-test for significance < 0.05.

Source Data

Extended Data Fig. 5 Resident alveolar type II cells, with GFP-labeled surfactant protein C (green) can be imaged inside the crystal ribcage.

(a) Stereomicroscope fitted with a NightSea GFP filter to visualize the SPC-GFP distribution at the whole organ scale. (b) Confocal microscopy of the SPC-GFP population at the lobe scale on the right side of the lung at 1.25x magnification. Vessels are labeled by injection of 50 µL of 10 mg/ml Evans blue dye in saline in vivo prior to extraction of the lungs. (c) Confocal microscopy at the 10x magnification shows single SPC-GFP+ cells in the lung distal parenchyma. (d) Single cell nuclei and SPC-GFP signal in cell cytoplasm visible at single cell resolution using a 60x water immersion lens.

Extended Data Fig. 6 Orthogonal views comparing the spatial distribution of nodular and infiltrative tumors, for three separate instances of each phenotype and solid phase nodular growth tumor increases imaging depth in sagittal image section using optical coherence tomography.

(a) Nodular tumors extended into the lung tissue to greater depths and bulge out of the pleural surface. (b) Infiltrative tumors are located more superficially on the pleural surface. (c) Alveolar function deeper than 100µm from the surface can be imaged through the tumor nodule up to a depth of 400µm from the surface, as the gas phase of the alveoli is replaced by the solid phase tumor, which reduces light scattering.

Extended Data Fig. 7 Single cell deformation in response to changing alveolar pressure.

(a) Cell-scale confocal microscopy of the lung inside the crystal ribcage with a FITC-albumin vascular lumen label showed the shadow of the cancer cell (b) which was imaged in a separate channel and merged to show cancer cells stretched in capillaries compared to arterioles and venules. (c) XY- views of the cancer cell at different depths of confocal imaging to show the cell is completely enclosed in the capillary. (d, e) Cells in capillaries quantified by the Feret ratio trended towards greater deformation in comparison to cells in larger arterioles-venules. Feret ratio quantified for n = 40 cells in capillaries, n = 30 cells in arteriole-venule at each alveolar pressure and data presented as mean ± SEM.

Source Data

Extended Data Fig. 8 Metastatic cancer cells disrupt vascular dye distribution in capillaries and the crystal ribcage used to visualize individual lung capillary walls and perfused lumens at high magnification (60x).

(a) Capillary scale confocal microscopy imaging of single cancer cells that disrupt the distribution of dye observed over timelapse imaging. (b) Vascular lumens were labeled with 10 mg/ml FITC-albumin dissolved in saline, injected intravenously prior to excision of the lungs for high-resolution confocal microscopy with a 60x objective.

Extended Data Fig. 9 MRP8+ Neutrophil cells co-localized with edema in lungs after 30 hours of lobar pneumonia injury.

Correlation of the edema label with neutrophil cells is imaged (a) using 1.25x, (b) 10x and (c) 60x objectives to confirm that all alveoli with neutrophils (green) are edematous (magenta). (d) The contralateral lobe was also imaged to show the absence of edema and neutrophils inside alveoli.

Extended Data Fig. 10 Lung adjacent organs such as the heart can also be imaged with the crystal ribcage.

Here using a laser-scanning confocal (a) the heart is imaged with a 1.25x objective to show (b) the overlapping lobes of the lung followed by (c) higher resolution images of the atrial surface and (d) muscle structures.

Supplementary information

Supplementary Information

Supplementary Notes, Methods and Figs. 1–14.

Reporting Summary

Supplementary Video 1

Functional ex vivo lung inside the crystal ribcage.

Supplementary Video 2

Whole lobe to single alveoli microscopy of lung under negative-pressure ventilation in the crystal ribcage.

Supplementary Video 3

Single alveoli microscopy of apex versus base in lung under negative-pressure ventilation in the crystal ribcage.

Supplementary Video 4

Crystal ribcage provides stable imaging surface for ventilated lung.

Supplementary Video 5

Immune cell migration in functional alveoli imaged in real-time through the crystal ribcage.

Supplementary Video 6

Apex to base image of lung with several metastatic tumors at alveolar resolution using OCT through the crystal ribcage.

Supplementary Video 7

Metastatic cancer cells remodel capillary function.

Supplementary Video 8

Real-time imaging of vascular transport at alveolar resolution in health and disease using crystal ribcage.

Supplementary Video 9

Neutrophil migration speed increases with higher vascular pressure.

Supplementary Video 10

Neutrophil extravasation and migration imaged through the crystal ribcage.

Supplementary Video 11

Local agarose blockage in the alveoli disrupts surrounding vascular dynamics.

Supplementary Video 12

Real-time imaging of spontaneous heart atrial pulsation in response to elevated pressure.

Supplementary Video 13

Air and perfusate leak from site of local excision in the lung.

Supplementary Video 14

Disrupted capillary flow in locally cauterized lung imaged through the crystal ribcage.

Source data

Source Data Figs. 1–5

Statistical source data.

Source Data Extended Data Figs. 2, 4 and 7

Statistical source data.

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Banerji, R., Grifno, G.N., Shi, L. et al. Crystal ribcage: a platform for probing real-time lung function at cellular resolution. Nat Methods 20, 1790–1801 (2023). https://doi.org/10.1038/s41592-023-02004-9

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