Supplementary Information for

Nonlinear optical endomicroscopy for label-free functional histology in vivo

Wenxuan Liang1, Gunnsteinn Hall1, Bernhard Messerschmidt2, Ming-Jun Li3, Xingde Li1*

1Department of Biomedical Engineering, Johns Hopkins University, Baltimore, Maryland 21205, USA2GRINTECH GmbH, Jena, Germany3Science and Technology Division, Corning Incorporated, Corning, New York 14831, USA*Corresponding author. E-mail: xingde@jhu.edu

This PDF includes:

Supplementary Note S1-S4

Supplementary Figure S1-S8

Supplementary Table S1-S2

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SUPPLEMNTARY NOTES

S1. System schematic and pulsewidth compensation

We adopted a grating pair and a dual-fiber strategy to manipulate and compensate the pulse

broadening due to both group delay dispersion (GDD) and spectrum narrowing induced by self-phase

modulation (SPM)1, 2 in the optical fibers. As illustrated in Fig. S1, femtosecond pulses from the

Ti:Sapphire laser (Chameleon Vision II, Coherent, Inc., Santa Clara, California, United States), with

an ~150-fs temporal pulse width, were first coupled into a piece of single-mode fiber (SMF, PM780-

HP, Thorlabs, Inc., Newton, New Jersey, United States), and then went through a grating pair for

dispersion compensation. The grating pair (600 lines∙mm-1, Wasatch Photonics, Inc., Durham, North

Carolina, United States) separation was tuned to compensate the total GDD in the SMF (~25 cm long)

and the customized DCF (~75 cm long). We found that that such dual-fiber configuration offered a

convenient way to both compensate dispersion and minimize SPM. And phantom experiments

demonstrated that such dual-fiber strategy could boost the two-photon excitation efficiency by at least

2X compared with the conventional single-fiber setup (manuscript in preparation).

The chirped excitation laser pulses were then launched into the DCF core and delivered to the

sample placed at the distal end of the endomicroscope. Emission photons, once epi-collected through

the achromatic miniature objective into the DCF inner clad, were guided back to the proximal end of

endomicroscope, where they were first separated from the excitation light by a long-pass dichroic

beamsplitter (FF665-Di02-25x36, Semrock, Inc., Rochester, New York, United States), and then

passed a short-pass optical filter (FF01-680/SP-25, Semrock) to further eliminate residual excitation

photons. The resultant signal photons could be spectrally filtered and detected by a photomultiplier

(PMT, H10771P-40, Hamamatsu Photonics K.K., Hamamatsu City, Shizuoka, Japan), as demonstrated

in Fig. S1. In the circumstances where dual-channel detection is needed (such as simultaneous

recording of NADH and FAD signals for redox ratio imaging), a second dichroic beam-splitter

(FF495-Di03-25x36, Semrock) could be used to further separate the emission photons spectrally

towards two separate PMTs. Appropriate optical filters could be placed in front of each PMT to further

define the spectral detection band. Optical filters used in this study included: 445/20 nm band-pass

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optical filter for SHG imaging (FF01-445/20-25, Semrock), 447/60 nm band-pass optical filter for

NADH signal (FF02-447/60-25, Semrock), and 496 nm long-pass optical filter for FAD signal (FF01-

496/LP-25, Semrock).

S2. Longitudinal focal shift characterization of the miniature objective

We adopted a confocal optical system to measure the longitudinal chromatic aberration of the

miniature objective3. To cover common excitation and emission wavelengths used in two-photon

microscopy, we combined light from three different lasers: one common blue laser (488 nm), one

common green laser (532 nm), and one supercontinuum laser (600-1000 nm, SuperK Extreme, NKT

Photonics, Birkerød, Denmark), as illustrated in Fig. S2. The combined light was coupled into one

input port (Port A) of an ultra-broadband fiber optic coupler (Gould Fiber Optics, Millersville,

Maryland, United States). Light coming out of the output port (Port C) was focused by the miniature

objective onto a silver mirror (with water immersion); then part of the back-reflected got coupled back

into the same single-mode fiber core (which serves as a confocal detection pinhole). Approximately

half of the collected back-reflected photons would exit another input port (Port B) and its spectrum

was measured by a spectrometer (BLUE-Wave VIS-25, StellarNet, Inc., Tampa, Florida, United

States). By scanning the mirror reflector longitudinally through the beam waist, a series of spectra of

the back-reflected light were recorded. From the stack of spectrum data, for each given wavelength of

interest, the reflector displacement corresponding to the peak back-reflection intensity was identified

and deemed as the optimal focal position (or sample-side working distance) for that wavelength.

This measurement method features three major advantages: 1) the usage of optical fiber mimics

exactly the actual application conditions of the miniature objective in the endomicroscope, which is

designed to focus divergent laser beam from the single-mode fiber core; 2) the confocality relationship

between the fiber-side focus (i.e. the fiber core in close vicinity of the fiber end surface) and the

sample-side focus of the miniature objective ensures efficient rejection of other wavelengths whose

focal planes fall farther away from the mirror surface, therefore guaranteeing sensitive and accurate

focal shift measurement; and 3) with the intensity variation of multiple wavelengths measured and

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recorded simultaneously, only a single mechanical scan of the reflector position is sufficient,

eliminating the need for position calibration between otherwise repeated mechanical scans.

S3. Collection efficiency simulation with the given inner-clad diameter and focal shift

The simulation framework for the endomicroscopy photon propagation has been described in detail

previously4. In essence, fluorescence photons were assumed to be generated inside the tissue at a

specified depth and distributed isotropically. Photon propagation inside scattering tissue was simulated

using the Monte Carlo Multilayered (MCML) approach5, where a water-immersion layer was placed

above the tissue layer. Photons exited the tissue surface with a direction vector v and position p, and

then passed through ray tracing calculations to evaluate whether they would be collected by the inner-

clad of the DCF. We studied the influence of DCF inner-clad diameter and the miniature objective

lens’ chromatic aberrations on the overall collection efficiency of two-photon emission from a tissue

phantom with a scattering mean free path of 92 µm (similar to tissue scattering property)6, and the

wavelength of emission (fluorescence) photons was assumed to be 500 nm.

As shown in Fig. S3, both the enlarged inner-clad diameter and the reduced focal shift of the

miniature objective are beneficial for collecting more scattered photons. Regarding the specific tissue

phantom considered here, our endomicroscope design with a DCF inner clad diameter of 185 µm and

a sample-side focal shift Δf = 10 µm (orange diamond in Fig. S3) can enhance the signal collection

efficiency by ~2.5~2.8X, when compared with the combination of a state-of-the-art GRIN objective

with a focal shift of Δf = 30 µm (GT-MO-080-018-810, GRINTECH GmbH, Jena, Germany) and a

commercial DCF (Nufern 5/130, NuFern, Inc., East Granby, Connecticut, United States) with an

inner-clad diameter of 130 µm (orange circle in Fig. S3).

S4. Redox ratio validation

To validate the measurements of the optical redox ratio (defined as FAD emission/(FAD emission +

NADH emission)), we calibrated our two-photon endomicroscope using NADH and FAD solutions.

Stock solutions (sealed from air) were made by disolving NADH (catalog number N8129, Sigma-

Aldrich Corporation, St. Louis, Missouri, United States) or FAD power (catalog number F6625,

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Sigma-Aldrich Corporation) in 1 mM Tris-HCl buffer with pH = 8.5. Then via serial dilution, standard

solutions of various concentrations were prepared for NADH (in ~0.1-3.2 mM range) and for FAD (in

~0.1-1.6 mM range).

First, the responses of our two-photon endomicroscope with dual-channel detection to these

standard NADH or FAD solutions were measured and plotted in the Fig. S7. The fluorescence

intensities were observed to scale linearly with the corresponding NADH or FAD concentrations.

Then the mixuture solutions were prepared with the FAD/NADH concentration ratio varying from

1/8 to 2/1 (or equivalently, the FAD concentration fraction varied from 1/9 to 2/3). Such ratio range

was selected based on the knowledge that typical cellular NADH concentration was estimated to be in

the ~100-400 μM range7-10 and FAD concentration in the ~10-150 μM range11-14. The NADH and FAD

fluorescence emission rates were measured from these mixture solutions, and then the optical redox

ratio, was calculated and plotted against the FAD concentration fraction in the solutions. As shown in

the Fig. S8, the optical redox ratio increases monotonically with the fraction of FAD. It is also noted

that the correlation of the measured optical redox ratio with the input FAD concentation fraction is

nonlinear, which could be caused by several factors including the difference between NADH and FAD

in two-photon action cross-section15 and system detection efficiency (e.g. spectra cut-off by optical

filters used for fluorescence detection), and potential cross-quenching of NADH fluorescence by

FAD16, 17 (which can happen both in solutions and in biological tissues).

Based on the above calibration curve, the optical redox ratio of ~0.9 we measured from the kidney

ischemia-reperfusion model (see Fig. 5 in main text) corresponds to a free FAD concentration fraction

of ~0.30. Factoring in the difference in two-photon action cross-section between protein-bound FAD

(~0.28×10-50 cm4∙s) and free-in-solution FAD (~0.071×10-50 cm4∙s)15, we can roughly estimate that the

overall FAD concentration fraction in the mouse kidney renal tubular cells ranges from ~0.075 to

~0.30; this aligns well with the previously reported cellular NADH and FAD concentration

estimations as mentioned above.

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SUPPLEMNTARY FIGURES

Figure S1 | Schematic illustration of the 2PF endomicroscopy system. See Note S1 for detailed

description of the system design. BP: band-pass optical filter. DCF: double-clad fiber. DM: dichroic

mirror. SP: short-pass optical filter. PMT: photomultiplier tube. SMF: single-mode fiber.

Figure S2 | Schematic of the experimental setup for longitudinal focal shift measurement .

Combined broadband light is first coupled into Port A, and then about 50% of the light exits Port C

and gets focused onto the mirror, which is scanned longitudinally through the focal region. Part of the

back-reflected light is coupled back into the single-mode core of Port C (which serves as a confocal

pinhole), and ~50% of the back-coupled light exits Port B and then is measured by a spectrometer.

CL: coupling lens; DM1: dichroic mirror (FT 580, Carl Zeiss AG, Oberkochen, Germany); DM2:

dichroic mirror (FT 510, Carl Zeiss AG, Oberkochen, Germany); OBJ: miniature objective; M: mirror.

See Note S2 for details about focal shift measurements.

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Figure S3 | Computed collection efficiency versus DCF inner clad diameter for various sample-

side focal shifts. The simulated tissue phantom used in computing collection efficiency has a

scattering mean free path of 92 µm and a scattering anisotropy (g) of 0.90. The imaging depth in the

tissue phantom was fixed at 100 µm for this simulation study. All collection efficiency values were

normalized with respect to the case of an ideal achromatic objective (corresponding to the black curve,

Δf = 0 μm) used in conjunction with an inner clad of a 300-µm diameter. Corresponding to the orange

diamond is our endomicroscope with a DCF inner clad diameter of 185 μm and a sample-side focal

shift Δf = 10 μm, while the orange circle indicates the combination of a state-of-the-art GRIN

objective (GT-MO-080-018-810, GRINTECH GmbH) and a commercial DCF (Nufern 5/130) with an

inner-clad diameter of 130 μm. See Note S3 for details of the simulation.

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Figure S4 | Single-frame endomicroscopy 2PF and SHG label-free structural imaging. The

images shown here are the corresponding single-frame version (frame acquisition time ~0.38 second)

of images shown in Fig. 3 of the main text, with the same excitation conditions: ~30 mW at 890 nm

(a-b); ~30 mW (c-d) at 750 nm, and ~40 mW (e-f) at 890 nm. Despite the discernible salt-and-pepper

noise, these single-frame images revealed the vast majority of essential structural details as seen in the

four- (or ten-) frame-averaged versions in Fig. 3. Scale bars, 10 µm.

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Figure S5 | Endomicroscopy 2PF redox imaging of a mouse kidney ischemia-reperfusion model

in vivo using the 300-µm WD miniature objective. The images shown here were acquired under

similar experimental conditions as those in Fig. 4 of the main text, except that the 300-µm WD

miniature objective was used instead. The essential structural characteristics and functional changes

revealed in Fig. 4 are also manifest here, demonstrating the similar performance of the two miniature

objectives. Top row (a-c): 2PF intensity images from the NADH detection channel (417-477 nm);

middle row (d-f): 2PF intensity images from the FAD detection channel (496-665 nm); bottom row (g-

i): 2PF intensity images color-coded by the measured optical redox ratio, defined as FAD/(FAD +

NADH), where more reddish (greenish) color represents a reduced (increased) redox ratio. The dark

round-to-elliptical spots scattered along the renal tubule wall (marked by dashed squares) correspond

to the nuclei of renal tubular cells, while the arrows (arrowheads) indicate the apical (basolateral) side

of the tubular cells. Each column corresponds to one specific time point: normal (left), 3 min 30 s

post-ischemia (center), and 3 min 05 s post reperfusion (right). All images are averaged over 5 raw

frames acquired with an incident power of ~33 mW at 750 nm, corresponding to an effective frame

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acquisition time of ~1.9 s. Scale bars, 10 µm.

Figure S6 | Single-frame endomicroscopy 2PF redox imaging of a mouse kidney ischemia-

reperfusion model in vivo. The images shown here were the corresponding single-frame version

(frame acquisition time ~0.38 second) of those images shown in Fig. 4 of the main text. Although

these non-averaged raw images appear noisier, similar structural and functional information are still

discernable. Top row (a-c): 2PF intensity images from the NADH detection channel (417-477 nm);

middle row (d-f): 2PF intensity images from the FAD detection channel (496-665 nm); bottom row (g-

i): 2PF intensity images color-coded by the measured optical redox ratio, defined as FAD/(FAD +

NADH), where more reddish (greenish) color represents to a reduced (increased) redox ratio. The dark

round-to-elliptical spots scattered along the renal tubule wall (shown in dashed squares) correspond to

the nuclei of renal tubular cells, while the arrows (arrowheads) indicate the apical (basolateral) side of

the tubular cells. All images here were acquired with an incident power of ~33 mW at 750 nm. Scale

bars, 10 µm.

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Figure S7 | Calibration curves of free NADH and free FAD solutions . Blue-diamond data points

represent actual fluorescence emission rate (in million photons/s) measured by our endomicroscope

with NAD detection band of 417-477 nm (a), and FAD detection band of 496-655 nm (b). Red

straight lines in both plots show the linear regression results, with coefficient of determination R 2 =

0.9986 for NADH, and R2 = 0.9971 for FAD.

Figure S8 | Calibration curve of optical redox ratio versus FAD concentration fraction in

solution. The free FAD/NADH concentration ratio is varied from 1/8 to 1/4, 1/2, 1/1, and 2/1 in the

calibration experiment, and the resulted FAD concentration fraction, defined as [FAD]/([FAD]+

[NADH]), is increased from 1/9 to 2/3, as indicated in the x-axis. The optical redox ratio, defined as

FAD emission/(FAD emission + NADH emission), was observed to grow monotonically with the FAD

concentration fraction. This calibration result validates the quantitative accuracy of the optical redox

ratio measured on the mouse kidney ischemia-reperfusion model with our two-photon

endomicroscope (see Note S4 for detailed analysis).

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SUPPLEMNTARY TABLES

Table S1. In-fiber background reduction by using the customized double-clad fiber (DCF)

Commercial DCFs for

comparison

Emission band of interest (nm)

Total (300-650) SHG (435-455) NADH (417-477) FAD (496-650)

SMM900 40.2 143 101 28.8NuFern 5/130 15.0 18.1 14.1 15.7

Note that within the SHG emission band, the ratio of background suppression is comparatively higher than the total background reduction ratio. The corresponding system sensitivity enhancement ratios can be readily calculated as the square roots of values shown in this table.

Table S2. Comparison of main parameters of the 200- and 300-μm WD miniature objectives

Miniature objectives

Working distance (µm) NA Diameter (mm)Color

correctionFiber-side (in air)

Sample-side (in water)

Fiber-side

Sample-side Optics Overall

Shorter WD 200 200 0.175 0.8 1.0 1.4 Yes

Longer WD 200 300 0.18 0.75 1.8 2.3 Yes

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