In recent years, there has been a tremendous number of new developments in optical microscopy techniques targeted at imaging different biological systems at a range of spatial and temporal scales, with each technique having its own intrinsic advantages and limitations. We focus on new developments in light sheet microscopy, also known as single plane or selective plane illumination microscopy.1,2
Light sheet illumination was developed at the start of the 20th century,3 and is having a resurgence in popularity after the characterization of fluorescent proteins4 and their rapid incorporation as genetically encoded visualization tools for biomedical microscopy applications.5 Light sheet microscopes were designed to illuminate specimens with a thin light sheet resulting in intrinsic optical sectioning for fluorescence microscopy.67.8.–9 Recent developments have significantly increased the resolution of this imaging method using multiple angle detection to achieve isotropic reconstruction of the specimen10,11 or via the implementation of lattice light sheet,12 the use of Bessel beams to appreciably reduce the thickness of the illuminated plane while keeping a large field of view, and a combination of Bessel beam excitation with two-photon illumination.13
Applications of light sheet microscopy are rapidly increasing, especially for imaging larger samples, such as zebrafish,14,15 fly embryos,9,16 C. elegans,17,18 and mouse organs or embryos.1920.–21 Due to the orthogonal illumination inherent to this technique, photobleaching and phototoxicity are significantly reduced compared with traditional laser scanning confocal microscopy,9,22 and single plane illumination permits the reduction of background fluorescence with camera acquisition of the entire imaging plane. This mode of detection significantly increases the image acquisition speed in comparison to point scanning systems, enabling three-dimensional (3-D) reconstructions of macroscopic samples and tissues in reasonable time periods and is especially suited for the imaging of live specimens.
Concurrently, sample clearing techniques have recently been developed to transform cumbersome opaque specimens into transparent samples for the optical imaging of highly scattering tissues, such as entire mouse brains or embryos. This property allows the observation of these biological specimens without relying on tissue sectioning and manual reconstruction, a process that can be time consuming and sometimes inaccurate. We utilize one of the simplest, cost effective, and rapid clearing techniques, iDISCO, which uses organic solvents to clear the tissues.23 However, dibenzyl ether (DBE), the final solvent of the protocol in which the sample is imaged, has a high-refractive index (), rendering most light sheet microscopes inoperable due to the change in refractive index between the objective and specimen. Some commercial solutions exist with specialized immersion objectives for water-based clearing techniques,24,25 but these objectives are relatively expensive and cannot be used for solvent clearing. These objectives, in addition to classic immersion objectives, cannot be used directly with such solvents without damaging the objective.
Here, we propose a new approach using a custom made holder which encases the sample and its medium in order to protect the objective (see Fig. 1). This overcomes limitations which would arise from direct contact solvent damage allowing us to mount standard objectives that can be used in conjunction with clearing solutions of any refractive index, in particular solvent-based clearing techniques. We implement our method using a simple modular passive light sheet design that incorporates an optomechanical decoupling of the excitation and detection objectives that enables us to image in organic solvents without a significant loss of resolution that would typically occur due to defocus from refractive index variations. We present the design and demonstrate applications of a system that is inexpensive, easy to build, and is scalable across sample type, size, and imaging medium.
Materials and Methods
SV129 mouse embryos imaged in this study were generously provided by Dr. Quenten Schwarz, University of Adelaide. Pregnant mice were euthanized at E11.5 and the embryos were collected, decapitated, and fixed in ice cold 4% paraformaldehyde (PFA) (Electron Microscopy Sciences, Hatfield, Pennsylvania) diluted in phosphate buffer saline (PBS) (141 mM NaCl, 2.7 mM KCl, 10 mM , and 1.5 mM ) overnight. For nerve preparations, adult C57BL/6 were raised according to the guidelines of the animal facility at the Montreal Neurological Institute. Adult mice were euthanized and nerves were collected and postfixed for 2 h in 4% PFA in PBS. Samples were washed in PBS and stored at 4°C prior to use.
Sample Preparation for Imaging
Whole mount fluorescent immunohistochemistry was performed according to a modified protocol.23 Samples were dehydrated using a gradient of methanol (50%, 75%, 100%) diluted in PBS, incubating for 1 h per methanol solution. Autofluorescence was quenched by incubating samples overnight in an ice cold mixture of 70% methanol (Fischer Scientific, Geel, Belgium), 20% dimethylsulfoxide (DMSO) (Sigma, St. Louis, Missouri), and 10% hydrogen peroxide (Fisher Scientific, Fair Lawn, New Jersey).
Samples were rehydrated with methanol solutions in the reverse order (75%, 50%, and PBS) and fixed embryos were blocked at room temperature for 12 h in 6% donkey serum (Sigma, St. Louis, Missouri), and 0.1% Triton-100 (Sigma, St. Louis, Missouri) in PBS. For visualization of the spinal cord commissural projections, embryos were stained for Tag-1 (AF4439, R&D Systems, Minneapolis, Minnesota) 1:200 in wash solution (0.6% donkey serum, 0.1% Triton X-100 PBS) for 48 h at 37°C. Samples were vigorously washed for 12 h and subsequently incubated in Alexa 568 conjugated antigoat donkey (IgG A11056, Invitrogen, Eugene, Oregon) (1:250 in wash solution) for 36 h at 37°C. Finally, samples were washed for 12 h with PBS and kept at 4°C until imaging.
Nerve samples were prepared following a similar protocol. After methanol dehydration and peroxide bleaching, rehydrated samples were blocked at room temperature for 12 h in 10% goat serum (Sigma, St. Louis, Missouri) and 0.2% Triton-100 in PBS. Sciatic nerves were costained with mouse anti-βIII-Tubulin (801201, BioLegend, San Diego, California, 1:33 dilution) and rabbit SCG10 (Stathmin-2) (NBP149461, Novus Biologicals, Oakville, Canada, 1:200 dilution), in nerve wash solution (3% goat serum, 0.2% Triton-X, heparin in PBS) for 5 to 7 days at 37°C. After extensive washing, samples were incubated with Alexa 488 conjugated antimouse IgG (Invitrogen, Eugene, Oregon) and Alexa 568 conjugated antirabbit IgG, at a dilution of 1:33 and 1:200, respectively, for 5 to 7 days at 37°C. Finally, nerves were washed for 12 h in nerve wash solution and 30 min in PBS before storage at 4°C until clearing.
Clearing was performed in accordance with a previously described protocol.26 Fluorescently stained embryos or nerves were first dehydrated using a tetrahydrofuran gradient (50%, 75%, and 100%) (Sigma, St. Louis, Missouri) at room temperature, for 1 h per solution. Samples were then immersed in DBE (Sigma, St. Louis, Missouri) for at least 30 min prior to imaging.
Human embryonic kidney 293T cells were cultured in Dulbecco’s modified Eagle’s medium, supplemented with 10% fetal bovine serum, 4 mM L-glutamine, penicillin, streptomycin, and 0.1 mM nonessential amino acids (Gibco, Carlsbad, California). Cells were maintained in a humidified 5% atmosphere at 37°C. Cells were grown for 2 days on coverslips before imaging. At 24 h postplating, plasmids encoding transmembrane proteins attached to EGFP and mCherry were transfected into the cells using LTX Lipofectamine with the Plus Reagent, as specified by the manufacturer (Life Technologies, Carlsbad, California). Imaging was performed in media consisting of Hanks’ balanced salt solution and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (Gibco, Carlsbad, California).
Interchangeable Sample Holder Design
Samples holders were designed (SOLIDWORKS, Waltham, Massachusetts) and cut out of acrylic sheets (1.5, 3, and 6 mm thick, Johnston Industrial Plastics, Montreal, QC, Canada) using a Roland MDX-40A CNC milling machine (Roland DGA Corporation, Irvine, California). Sample holders are displayed below in Fig. 1.
For encapsulation of samples containing DBE, two parts are milled, aligned, and assembled using acetone (ACP, Montreal, QC, Canada). After the sample holder has dried (), a coverslip (#1) is attached to the front side using clear nail polish, and left to dry overnight. The attached coverslip is then shaped using a diamond burr to fit the front side of the holder.
Multimodal Light Sheet Setup with Dynamic Decoupling Between Excitation and Detection
We designed and built a highly modular system around a commercial microscope base creating a user-friendly light sheet microscopy system. We used a Nikon FN-1 upright microscope base (Nikon Canada Inc., Mississauga, ON, Canada), where the back left port of the NI-TT Quadrocular tilting tube on top of the microscope was used for detection. The microscope was equipped with halogen and mercury lamps for rapid sample visualization and positioning. An overview of the experimental setup for this microscope is shown below Fig. 2.
Imaging was performed with three different detection objectives for a variety of resolutions and fields of view; CFI Plan Apo Lambda (0.45 NA, Nikon), CFI75 Apo LWD (1.1 NA, Nikon), and CFO Apo (0.8 NA, Nikon). A long travel objective piezoscanner (P-725.4CD, Physik Instrumente L.P., Auburn, Massachusetts) allowed rapid and reproducible movement of the objective lens. There are two main light pathways; 100% binocular and 100% camera port, with four different tube lenses (, , , ). The whole setup was assembled on an optical breadboard table (Newport Corporation, Irvine, California).
For excitation, we designed a laser box with five OBIS diode lasers (405, 445, 488, 561, 604 nm, Coherent Inc., Santa Clara, California) to cover excitation wavelengths for the majority of fluorophores, with two fibered and one free space outputs. All five laser lines were combined using reflective and dichroic mirrors, and directed to one of the three outputs using switchable mirrors. For light sheet use, the laser lines were focused onto a fiber (OZ Optics Ltd., Ottawa, ON, Canada) and the fiber output was directed to an excitation arm, where light was collimated using a lens and then shaped into a sheet of light via a cylindrical lens placed before a CFO Apo (0.8 NA, Nikon) or CFI Plan Fluor (0.3 NA, Nikon) excitation objective.17 The excitation arms are assembled on a magnetic base to be easily interchangeable and are mounted on two linear stages to adjust the plane of light.
The sample holder module was placed on the right side of the microscope, opposite to the excitation arm, specially designed to fit the range of sample holders, Fig. 1, within the working distance of the appropriate objective. It was assembled on two sets of , , and linear motors: one manual set for rapid sample positioning at the point of convergence between the excitation and detection objectives, and a second automated set for reproducible movement within the sample and sample stitching for larger fields of view. Additionally, two linear ( and ) piezoactuators were added for fast 3-D imaging at nm step resolution. The system can also be modified for cuvette samples as shown in Fig. 3.
The entire setup, including excitation, detection, and all motorized parts, was controlled using LabVIEW and an NI-DAQ acquisition card (NI-PCIe-6353, National Instruments, Toronto, ON, Canada). The lasers, camera, and piezocontrollers are triggered using a digital or analog signal from the NI-DAQ for synchronization with the internal clock. Emitted photons are collected by the objective, filtered through a quad-band fluorescent cube (Chroma Technology Corp., Bellows Falls, Vermont), and detected on a Andor iXon Ultra 888 EMCCD camera (SnowHouse Solutions Inc., Lac-Beauport, Quebec).
A highly detailed supplementary file is available to any interested researcher via simple e-mail request. All parts are listed in this document with all CAD drawings and procedures for alignment and programming.
Results and Discussion
Combining Multiple Light Sheet Modes into One System
An important goal in microscope development is to design a way to combine many different modalities into one system. In the specific case of light sheet microscopy, the objective working distance is a limitation, as it needs to accommodate the sample size while maintaining a large numerical aperture for detection purposes. In the generation of a light sheet, only a section of the sheet can be considered homogenous, restricting the field of view. Furthermore, better resolution in the direction can only be achieved by decreasing the field of view, e.g., a light sheet thickness of typically has a field of view of , whereas to attain a thinner sheet, the field of view will be reduced to . To our knowledge, outside of high-end solutions such as isotropic multiview10,11 or lattice light sheet12 that can allow, respectively, a greater or both better and field of view, there is no solution to this trade-off. However, we introduce a very low cost simple adaptive solution which can be used on any sample size and desired resolution.
In our system, three different excitation arms are made easily interchangeable to fit the user’s needs, see Fig. 3. The first excitation arm employs the objective, Figs. 3(a) and 3(b), to provide a relatively large field of view and a sizeable working distance of 16 mm, which can be used with any detection objective. The system can be easily adapted for imaging in a cuvette with a coverslip for side illumination, or for the homemade sample holders, shown in Fig. 1, to envelope specimens and protect objectives when used with organic solvents. However, this size excitation objective generates a sheet of .
The second excitation arm utilizes a objective [Fig. 3(c)] with a working distance of 3.5 mm, which can only be used with an identical detection objective. A cuvette is needed for the immersion of both objectives and a sample holder can be used to encase cleared samples in organic solvent, while protecting both objectives. The light sheet produced from this objective is thick, with a relatively small field of view of .
Decoupling Excitation and Detection Planes
Here, we present a straightforward solution to compensate for changes in refractive index that can be easily implemented on the majority of light sheet systems. Clearing techniques are required for the imaging of relatively large samples with fluorescence microscopy. However, these methods often employ solvents with optical indices significantly different from the immersion liquid, i.e., water. Our implementation aims to overcome this limitation and can be applied to liquids of any refractive index.
When imaging in media with a different refractive index than that required for the detection objective, the detection cone [dotted line in Fig. 4(a)] is modified [solid line in Fig. 4(a)]. The numerical aperture and working distance for detection will be altered depending on the distance needed to travel through each medium to reach the objective. This can become problematic when scanning the sample along the detection axis. The defocusing effect of an index mismatch is illustrated in Figs. 4(b) and 4(c). As light sheet microscopy decouples excitation and detection into two objectives, when moving the sample the excitation and detection planes will quickly dissociate from each other. To compensate for the discrepancy in working distance, we implement a facile solution by relocating the objective Fig. 4(d), which recovers the overlap needed between the excitation sheet and detection plane.
Rapid Correction of the Imaging Plane
For correction of the imaging plane in fast acquisition mode (20 to ), the objective needs to be rapidly repositioned for each frame. In our system, we implemented this via a piezoscanner for the objective to synchronize with the piezoactuator scanning the sample in the light sheet. Synchronization was made possible by simultaneously sending an analog signal via NI-DAQ to both piezosystems and a digital signal to the lasers and the camera.
For imaging in any refractive index medium, the compensation displacement of the objective is configured by the user determining the correct position for the piezoactuator for the top and the bottom of the stack in order to obtain a focused image.
When these parameters are known for a given imaging medium, the user can utilize the same displacement or redefine the parameters. During our experiments, a typical correction for the objective in DBE is for a stack and for the objective. After user setup, the software simply divides the entire compensation displacement by number of planes in the stack in order to apply the compensation during the imaging of each plane
By using these live corrections, users can image in any refractive index environment and we have demonstrated this in DBE, Fig. 5. We show the detrimental impact on imaging without the piezoelectric correction, exhibited after a small displacement of in Fig. 5(a), and we present the acquisition with the correction from our technique in Fig. 5(b), revealing structures that were previously obscured. Users can acquire large stacks without a loss of information or the need to periodically refocus. Video 1 highlights the same corrective implementation for 3-D imaging of a mouse embryo.
Acquisition of an Entire Specimen
When using the excitation arm equipped with the objective, the field of view is considered to be (i.e., homogeneous part of the light sheet) by (camera field of view) and the maximum movement of the piezoelectric actuator with respect to the stack is . In order to record the entire specimen (i.e., 12.5-day mouse embryo, 4 ), the acquisition needs to take place over multiple fields of view. For this, we use three motorized stages to take stacks at different positions, crop each stack to keep the thinnest homogeneous part of the light sheet, and stitch the stacks together as shown in Fig. 6.
Automated data treatment for the acquisition of the entire specimen was performed using the Fiji stitching plugin27 and 3-D rendered using Imaris (Bitplane, South Windsor, Connecticut). For the sample presented in Figs. 5 and 6, acquisition time was around . The acquisition time varies depending on the integration time and camera mode.
Spinal cord commissural neurons are an important model system for axon guidance in mammals. This population of neurons projects their axons radially toward the ventral midline, where they cross to the opposite hemicord and change their growth direction to the higher brain regions.28 These directional changes make it difficult to quantify the different possible phenotypes for these axons. One limitation in the field is the need to physically slice the samples and perform immunostaining for the neuronal populations of interest, but this has recently been overcome by the development of clearing techniques.23,26,29,30
As hundreds of thousands of commissural neurons project in parallel, only an imaging system capable of resolving very fine structures would allow us to study the neuronal projections. This can be achieved by reducing the light scattering of the sample through clearing using a range of different chemicals that act by equilibrating refractive indices through the tissue.31 Each clearing method uses a mounting medium with a specific refractive index31 and this often poses a problem for microscope objectives, which are usually designed for a single refractive index.
As can be observed in Fig. 5(a), the structure of thick spinal cord commissural axons ( to ) is lost as we advance deeper in the DBE-immersed tissue. By mechanically correcting the focal plane deviation caused by DBE, we are able to visualize the crossing fibers in the midline, Fig. 5(c). To our knowledge, this is one of the sharpest images of the developing commissural spinal cord tracts. Using our method for focus correction, we were able to image all of the spinal cord commissural neurons using the general marker Tag-1, at a similar resolution to what has been previously published with neuronal subpopulation markers.29 Thus, the imaging improvement brought by mechanical focal correction allows for a detailed inspection of spinal cord phenotypes without incurring a cell population bias in order to resolve the axons.
In addition, clearing techniques enable the observation of whole mouse embryos without tissue sectioning, Fig. 6. As the neurons in the spinal cord differentiate in waves, in which the cells at the brachial level mature earlier than the caudally located neurons,32 it is advantageous to be able to resolve the complete structure. In comparison to commercially available light sheet microscopes, mechanical focal correction can be used to reach the same resolution without specific sample preparation constraints and index compatibility.
The benefits of our correction method are readily apparent when applied to nerve imaging. Understanding how nerves develop and regenerate is a major challenge for the neuroscience field. The current gold standard method for this analysis is the manual sectioning of nerves in transverse or longitudinal sections and subsequent reconstruction. For a rodent nerve, that usually implies combining images of hundreds of slices from single nerves in order to find growing fibers. In such cases, one nerve can have a diameter of 0.5 mm and length of several centimeters. As the sectioning process transposes a 3-D system to two-dimensional fragments, it is susceptible to many interpretation mistakes, including the confusion of spared and regenerating fibers.33
While multiphoton and point scanning confocal microscopy have been used together with tissue clearing, these techniques are time consuming and can give rise to a significant bleaching problem.34 These drawbacks can be overcome with spinning disk microscopy if pinhole crosstalk did not pose a challenge for scanning deep tissues.35 To our knowledge, some of the best results for the imaging of regenerating nerves in whole mount preparations have been obtained by light sheet imaging of cleared tissue using organic solvents.36,37 Here, we show that our system is capable of imaging both DBE- immersed sciatic nerves, which are important models for the study of the peripheral and central nervous systems, respectively. We can observe the expression of neuronal markers, such as β-III-tubulin, to follow axon morphology across an entire nerve, as shown in Fig. 7. Regenerating fibers can be identified by Stathmin-2 expression, a marker of growing and sensory axons,38 also shown in Fig. 7. Our system could enable us to count and trace growing and regenerating fibers throughout the extension of the nerves.
Altogether, light sheet microscopy consists of a practical and accessible method to image whole mount preparations. Combined with paradigms of nerve lesion, this system can be used to accurately assess nerve regrowth in future studies.
Imaging of Live Single Cells
Equipping the light sheet microscope with objectives for excitation and detection, alongside the 45 deg sample holder, Fig. 1(b), single cell imaging was achieved at a 3-D acquisition timescale of ms. However, using this objective, the field of view was limited to 20 to . The piezoelectric actuators were completely synchronized, as one was used to scan the sample in the direction, while the other simultaneously moves the sample along the excitation axis in order to center the sample at the thinnest part of the light sheet for the best resolution. Images of single cells are shown in Fig. 8.
Implementation of Further Corrections
For compensation of the defocusing effect when stitching large specimens, the excitation arms can be further motorized along the excitation axis, analogous to our solution employed to correct for excitation in high-refractive index media. In our system, the light sheet displacement along the excitation axis was almost negligible due to the numerical aperture and working distance of the objective used, therefore, we did not need to motorize this part. However, other objectives would need this correction. The cropping of data to include only sampling from the thinnest part of the light sheet enables the best resolution possible.
Aberrations due to material scattering and absorption (i.e., stripes) can be avoided by the use of two light sheets, generated by two objectives on different sides of the sample,8 by pivoting the cylindrical lens,8 by implementing a rotational stage,7,19,20 or by software treatment.39 Currently, we are adding a third excitation arm, Fig. 3(d), to our modular system to enable the generation of a lattice light sheet, for improved resolution in while keeping a reasonable field of view.12
In conclusion, we have introduced and described a highly modular light sheet microscope design equipped with different excitation arms for a variety of imaging samples and conditions and a wide array of sample chambers/holders. Our goal is to enable interested readers to build their own light sheet systems, even with minimal knowledge of optics and programming, where the system is entirely flexible to image at different resolutions.
As described recently in a comprehensive review,40 the best clearing techniques are challenging for microscopy due to the large indices of refraction for the solutions typically used. The system presented here enables users to work in any refractive index imaging medium by optomechanically decoupling the excitation and detection movement through a straightforward implementation of two synchronized piezoelectric actuators. Through our solution, we introduce live compensation along the detection axis during stack acquisition for every refractive index, enabling high resolution 3-D imaging of specimens cleared using a variety of techniques. To our knowledge, this is the first light sheet microscope to enable such versatility of imaging in media with varying refractive indices.
We acknowledge the Integrative Quantitative Biology Initiative for support. 3-D rendering for this work was performed in the McGill University Life Sciences Complex Advanced BioImaging Facility. SV129 mouse embryos were kindly provided by Quenten Schwarz (University of Adelaide). We acknowledge Claire Chardes for her initial help on light sheet microscope design. Purchase of equipment was made possible with funding from the Canadian Foundation for Innovation. This work was supported by operating grants from the Natural Sciences and Engineering Research Council of Canada to P.W.W. and from the Canadian Institute of Health Research and the Multiple Sclerosis Society of Canada to A.E.F. V.R. acknowledges financial support from the NSERC-CREATE Bionanomachines Program. R.A., M.-P.G., and A.A.G. are supported by the NSERC-CREATE Neuroengineering Program. R.A. also acknowledges financial support from CIHR/ Vanier CGS. A.K. is a recipient of a Fonds de Recherche du Quebec–Nature et Technologies doctoral scholarship.
Vincent Rouger received his BSc degree in physics and chemistry from Burgundy University in 2005 and his PhD in immunology from Aix Marseille University at the Centre d’Immunologie de Marseille Luminy in 2013. Currently, he is working as a postdoctoral fellow in Professor Wiseman’s laboratory in the Chemistry Department at McGill University.
Ricardo Alchini received his BSc degree in biological sciences from the Federal University of Santa Catarina in 2008 and his MSc degree in frontier biosciences from Osaka University in 2011. Currently, he is a PhD candidate in Professor Fournier’s laboratory in the Montreal Neurological Institute at McGill University.
Alexei Kazarine received his BSc (Hons) degree in chemistry at McGill University in 2012 and went on to complete his MSc degree in chemistry, specializing in centrifugal microfluidics applied to analytical chemistry in 2014. Currently, he is a PhD candidate in the Department of Chemistry in Professor Wiseman’s laboratory, using nonlinear microscopy to investigate biological tissues.
Angelica A. Gopal received her MSci (Hons) degree in chemistry with an international study year from the University of Nottingham in 2011, specializing in physical chemistry. She is completing her PhD in the Department of Chemistry in Professor Wiseman’s laboratory, using image analysis techniques applied to biological systems.
Marie-Pier Girouard received her BSc degree in biochemistry from the Université de Sherbrooke in 2010. Currently, she is pursuing her PhD studies in the integrated program of neuroscience at McGill University in the laboratory of Professor Alyson Fournier.
Alyson E. Fournier is a full professor in neurology and neurosurgery at McGill University. Her research lab is at the Montreal Neurological Institute and studies molecular and cellular responses to central nervous system injury to develop therapeutic strategies to promote repair.
Paul W. Wiseman received his BSc degree from St. Francis Xavier University in 1989 and his PhD from the University of Western Ontario in 1995. In 2001, he started as an assistant professor jointly appointed in the Departments of Chemistry and Physics at McGill University, was promoted to associate professor in both departments in 2007, and to full professor in 2013. His research focuses on biophysics measurements of protein interactions and transport in living cells and neurons.