This work was supported by NSF DBI-1707312 Cornell NeuroNex Hub and NIH 1U01NS103516.

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The authors declare no competing interests.

This protocol explains step-by-step procedures for setting up 3P imaging with a commercial microscope and laser source. Compared to 2PM, 3PM has an advantage in applications requiring optical access in the deeper regions such as in the mouse brain hippocampus. Although 3PM is mostly used in neuroscience, 3PM can be potentially applied in other tissues such as lymph nodes, bones, and tumors for deep-tissue observation.
It is important to verify that the imaging system performs at close to the shot-noise limit, which ensures that the detection and data acquisition electronics contribute negligible noise to the image after the PMTs. The uncertainty in the number of photons detected is fundamentally limited by photon shot noise. Shot-noise limited performance can be achieved in a typical multiphoton microscope using a high-gain photodetector (e.g., a PMT). Photon shot noise follows a Poisson statistical distribution, wherein the standard deviation of the distribution is equal to the square root of the mean of the distribution. To verify the shot-noise limited performance, follow step 1.14 in the protocol section.
To avoid light attenuation by H2O, using D2O for immersion is helpful, particularly for ~1,700 nm excitation. When D2O is used, it is essential to refresh D2O every ~10 min or use a large volume of D2O to avoid D2O/H2O exchange during imaging. One can also seal the D2O from the room environment3. If a long working distance (WD) objective lens (e.g., WD at 4 mm or longer) is used for imaging, the immersion liquid thickness can exceed 2-3 mm. The increased thickness makes H2O absorption non-negligible even at ~1,300 nm21. Therefore, D2O may be necessary even for 1,300 nm 3PM when using a long WD objective lens.
As the 3P fluorescence intensity depends on the cube of the excitation pulse energy at the focus (Eq. (1)), setting the appropriate laser power is particularly important to obtain adequate 3P fluorescence signals while avoiding thermal and nonlinear damage in living tissues. The average laser power should be kept below the thermal damage threshold. In the mouse brain, for example, to avoid thermal tissue damage, the average power on the mouse brain surface should be kept at or below ~100 mW for ~1,300 nm excitation at a depth of 1 mm and with a field-of-view (FOV) of 230 &#181;m x 230 &#181;m21. Similarly, the average power at ~1,700 nm should be kept at or below ~50 mW at ~1 mm depth and a FOV of ~230 &#181;m x 230 &#181;m (unpublished data). Further, to avoid excitation saturation and potential nonlinear damage, the excitation pulse energy should be kept at <img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />2 nJ and <img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />3 nJ for ~1,300 nm and ~1,700 nm excitation, respectively30.
Due to light absorption and scattering in tissues, the pulse energy at focus is attenuated to 1/e (~37%) after penetration of tissues by 1 EAL. The EAL varies in different tissues and with the excitation wavelengths, e.g., in the neocortex of the mouse brain, the EAL is ~300 &#181;m and ~400 &#181;m at ~1,300 nm and ~1,700 nm, respectively3,29&#160;(Figure 5). Therefore, to keep the same pulse energy at focus (e.g., 1 nJ/pulse) at a depth of n EALs, the surface pulse energy needs to be multiplied by 1 nJ &#215; en. For fast imaging of structural and functional dynamics, an excitation laser with a high repetition rate (at 1 MHz or higher) is desirable to achieve a high frame rate5,6,7,10. However, the pulse energy requirement and the average laser power limit constrain the applicable repetition rate.
For example, when we image a moderately deep region at 4 EALs (i.e., ~1.2 mm in the mouse cortex with 1,300 nm excitation), ~55 nJ/pulse at the surface is required to keep 1 nJ/pulse at focus. When the average power limitation is 100 mW, we can apply a ~2 MHz laser repetition rate. However, to image deeper at a depth of 7 EALs, ~1,100 nJ/pulse is required at the surface to maintain 1 nJ/pulse at focus. Assuming the maximum average power is 100 mW to avoid thermal damage, the laser repetition rate should be reduced to 0.1 MHz to achieve a 1,100 nJ/pulse at the surface. Table 1&#160;summarizes typical imaging conditions in the mouse brain cortex. Note that the imaging depths in Table 1&#160;assume that the EAL is uniform in the entire mouse cortex.
Moreover, because of the laser power limitation in deep-tissue 3PM, a trade-off exists between the frame rate and the image pixel size, which is particularly important for functional imaging such as calcium imaging. The maximum available laser repetition rate is decided at each depth based on the required pulse energy at focus and the applicable average laser power as discussed above, e.g., 2 MHz at a depth equivalent to ~4 EALs with 1,300 nm excitation. In general, imaging requires at least one pulse per pixel. Accordingly, the minimum available pixel dwell time is determined by the laser repetition rate, e.g., 0.5 &#181;s/pixel with 2 MHz excitation.
To keep the high spatial resolution (~1 &#181;m in lateral) in 3P images, it is ideal to set 1 pixel to an area of ~1 &#181;m2, e.g., 256 x 256 pixels for a FOV of 250 x 250 &#181;m2. Hence, to perform fast imaging with a considerably large FOV (e.g., 250 x 250 &#181;m2 with 256 x 256 pixels), 0.5 MHz, 1 MHz, and 2 MHz pulse repetition rates give theoretical maximum frame rates of ~7.6, ~15, and ~30 frames/s, respectively. Likewise, the optimization of the laser repetition rate is essential, depending on the target depth, scan speed, and FOV, to apply adequate pulse energy under the thermal damage threshold. To increase the imaging speed, an adaptive excitation source can be used to concentrate all the excitation pulses on the neurons (i.e., regions of interest) by delivering laser pulses on demand to the neurons31.
3PM is advantageous when compared to 2PM in deep imaging within living tissues and through highly scattering media such as a skull, bones, and the white matter layer (i.e., the external capsule) of the mouse brain. The longer EAL and the higher-order nonlinear excitation of 3PE benefits deep-tissue imaging. For example, to image GCaMP6 in the mouse cortex, 2P&#160;fluorescence signal with 920 nm excitation is higher than 3P&#160;fluorescence signal with 1,300 nm excitation in shallow regions at <img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />690 &#181;m (i.e., ~2.3 EALs at 1,300 nm)21. However, due to the longer EAL at 1,300 nm compared to 920 nm, 3PE gives stronger fluorescence than 2P excitation (2PE) at a depth of ~690 &#181;m and deeper21. This depth is defined as &#39;signal crossover depth,&#39; at which the fluorescence signal strengths of 2PE and 3PE are identical with the same repetition rate and the same maximum allowable average powers21. The signal crossover depth depends on the excitation wavelengths for 2PE and 3PE and the fluorophore.
In practice, 920 nm excitation allows higher average laser power than 1,300 nm excitation due to less water absorption. However, the higher average power of 2PE would push the signal crossover depth only by 0.9 EALs4. In addition, when the sample is densely labeled, 3PE has the additional advantage of much higher SBR. Therefore, even before reaching the signal crossover length, 3PM can be better for imaging than 2PM. For example, when imaging the mouse brain vasculature, which has a volume fraction (i.e., labeling density) of ~2%, 1,300 nm 3PM with 100 mW excitation power outperforms 920 nm 2PM with 200 mW excitation power at a depth of ~700 &#181;m for fluorescein.
3PM also has an advantage when imaging through a thin but highly scattering layer that may distort the point-spread function of the excitation beam and generate a defocus background4. For example, through the intact skull of the mouse brain, 2PM images suffer from the defocus background even at the shallow depth of &#60;100 &#181;m from the brain surface13. A similar defocus background was observed in 2PM with 1,280 nm excitation through the white matter in the mouse brain32. Therefore, when tissues are imaged through turbid layers, 3PM is preferable to 2PM for high-contrast imaging regardless of the labeling density.
We recently reported a beads phantom and theoretical analysis showing that the imaging depth limit of 3PM is over 8 EALs33; 8 EALs are equivalent to ~3 mm with ~1,700 nm excitation in the mouse cortex. However, the currently available laser does not have enough pulse energy to achieve 8 EALs in the mouse brain. Further development of stronger lasers will push the current imaging depth limit of 3PM.

In life science, multiphoton microscopy (MPM) techniques, such as 2PM and 3PM, have been powerful tools for deep in vivo imaging with high spatiotemporal resolution and high contrast in scattering tissues. Additionally, these methods cause less photobleaching when compared to one-photon confocal microscopy1,2,3,4. 3PM is advantageous for deeper-tissue imaging when compared to 2PM due to two major features : (i) employment of longer wavelength excitation (~1,300 nm or ~1,700 nm) reduces tissue scattering, and (ii) the higher-order excitation process (i.e., fluorescence signal depends on the cube of the excitation power in 3PM instead of the square of the excitation power in 2PM) that suppresses the unwanted background fluorescence3. Consequently, 3PM enables high-contrast imaging at deeper regions in living tissues such as the hippocampus in an intact adult mouse brain3,5,6,7,8,9,10,11&#160;and the entire forebrain of adult zebrafish12, including Ca2+ activity recording and multicolor observations. Moreover, high-contrast images have been obtained with 3PM through the intact skulls of mouse and adult zebrafish12,13.
Today, excitation laser sources suitable for 3P excitation (3PE) at ~1,300 and ~1,700 nm are commercially available. As the laser scanning system is essentially the same for 2PM and 3PM, converting an existing a 2P&#160;setup to a 3P setup is possible in biology laboratories with the installation of a commercially available laser for 3PE. The 3P fluorescence signal is dependent on the laser power, pulse duration, laser repetition rate, and numerical aperture (NA) of the objective lens. Assuming a diffraction-limited focus (i.e., the back aperture of the objective lens is overfilled by the excitation beam), Eq (1) describes the time-averaged fluorescence photon flux from the focal volume resulting from 3PE.
<img alt="Equation 1" src="/files/ftp_upload/63213/63213eq01v3.jpg" /> (1)
Where f is the laser repetition rate, &#964; is the laser pulse duration (full width at half maximum), &#981; is the system collection efficiency, &#951; is the fluorescence quantum efficiency, &#963;3&#160;is the 3P absorption cross-section, C is the fluorophore concentration, n0&#160;is the reflective index of the sample medium (e.g., water), &#955; is the excitation wavelength in vacuum, NA is the numerical aperture of the objective lens, a3&#160;is the spatial integration constant of the focal volume, <img alt="Equation 1" src="/files/ftp_upload/63213/62313eq02.jpg" /> is the time-averaged excitation photon flux (photons/s) under the objective lens, z is the imaged depth, and EAL is the effective attenuation length14. Here we have assumed that the EAL (typically &#62; 100 &#181;m) is much greater than the axial resolution of the microscope (typically &#60; 10 &#181;m). Under paraxial approximation, a3&#160;is equal to 28.114. gp(3) is the 3rd-order temporal coherence of the excitation source, and gp(3) is 0.41 and 0.51 for hyperbolic-secant-squared pulses and gaussian pulses, respectively. The collection efficiency &#981; can be estimated by considering the fluorescence collection by the objective lens, transmittance of the objective lens, the reflectivity of the dichroic mirror, the transmittance of the filters, and the detection efficiency of the detector (e.g., photomultiplier tube, or PMT). As the 3P fluorescence intensity is highly dependent on various parameters, optimization of the 3P setup is required to maximize the 3P fluorescence signals.
This protocol illustrates the optimization process of a typical 3P setup, which will be useful particularly for biology laboratories that have a 2P setup and plan to expand its capability to 3P imaging or to maintain their commercial 3P setup at optimal performance. This video article also demonstrates deep-tissue 3P imaging in living animal brains. The first section addresses the optimization of a typical 3P setup with a commercially available laser source and multiphoton microscope. The second and third sections describe zebrafish and mouse preparation, respectively, for 3PM of neuronal structures and activities. The mouse craniotomy surgery has been previously reported in protocol papers as well15,16,17. The fourth section demonstrates intravital 3P imaging in zebrafish and mouse brains.

<table><tbody><tr><td>5% Povidone-iodine</td><td>Amazon</td><td>NDC 67818-155-32</td><td>Aceptical cleaning of surgical areas</td></tr><tr><td>70% Ethanol</td><td>Thermo Fisher Scientific</td><td>CAS 64-17-5</td><td>Aceptical cleaning of surgical areas</td></tr><tr><td>Agarose</td><td>Sigma</td><td>A4718-256</td><td>Preparing zebrafish chamber</td></tr><tr><td>Atropine</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Bergamo II</td><td>Thorlabs</td><td></td><td>Multiphoton Imaging Microscope</td></tr><tr><td>Bupivacaine</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Dexamethasone</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Donut shape glass (ID4.5, OD6.5)</td><td>Potomac Photonics</td><td></td><td>Cover glass used for craniotomy</td></tr><tr><td>eye ointment (or topical ophthalmic ointment)</td><td>Puralube Vet Ointment</td><td>NDC 17033-211-38</td><td>Used as a lubricant to prevent irritation or to relieve dryness of the eye during surgery and anesthesia</td></tr><tr><td>GaAsP Amplified PMT</td><td>Thorlabs</td><td>PMT2100</td><td>PMT detector</td></tr><tr><td>Glucose</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Glycopyrrolate</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Heater (800 W)</td><td>Finnex</td><td></td><td>Aquarium heater for zebrafish water)</td></tr><tr><td>Isoflurane USP 250 mL</td><td>Piramal</td><td>NDC 66794-0013-25</td><td>For anesthesia of mice</td></tr><tr><td>Ketoprofen</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Kimwipes</td><td>Kimtech</td><td></td><td>Laboratory tissue for preparing zebrafish</td></tr><tr><td>Nanofil syringe (10 micrometer) with 36 G needle</td><td>WPI</td><td>NANOFIL + NF36BV</td><td>Syringe and needle for injection of pancuronium bromide</td></tr><tr><td>Optical Adhesive</td><td>Norland</td><td>NOA 68</td><td>To stick round coverslip and donut shape glass together.</td></tr><tr><td>Pancuronium Bromide</td><td>Cornell Veterinary Care</td><td></td><td></td></tr><tr><td>Peristaltic Pump</td><td>Elemental Science</td><td>ESI MP2</td><td>Water pump for zebrafish setup</td></tr><tr><td>Polyethylene tubing (I.D. 0.58 mm., O.D. 0.965 mm.)</td><td>Elemental Science</td><td>MP2 pump tubing</td><td>Tubing that goes in the mouth of the zebrafish</td></tr><tr><td>Round Cover Slip German Glass #1.5, 5 mm</td><td>Electron Microscopy Sciences</td><td>7229605</td><td>Cover glass used for craniotomy</td></tr><tr><td>Spirit-NOPA</td><td>Spectra Physics</td><td></td><td>Tunable Optical Parametric Amplifier</td></tr><tr><td>SR400</td><td>Stanford Research Systems</td><td>SR400</td><td>Photon counter</td></tr><tr><td>Standard Photodiode Power Sensor</td><td>Thorlabs</td><td>S122C</td><td>Power detector</td></tr><tr><td>Sterilized phosphate buffered saline (PBS)</td><td>Millipore Sigma</td><td>SKU 806552-500ml</td><td>Used during mouse brain surgery</td></tr><tr><td>Surgical drape</td><td>Dynarex disposable towel drape</td><td>4410</td><td>For aceptical mouse surgery</td></tr><tr><td>Thin strip boxing wax</td><td>Corning Rubber Co., Inc.</td><td></td><td>Holding tubing in place in zebrafish chamber</td></tr><tr><td>ThorImage</td><td>Thorlabs</td><td></td><td>Image acquisition software</td></tr><tr><td>Tricaine (Ethyl-m-aminobenzoate methanesulfonate salt)</td><td>MP</td><td>103106</td><td>Zebrafish anesthesia and euthanasia</td></tr><tr><td>Tygon tubing (I.D. 1/16 in., O.D. 1/8 in.)</td><td>Tygon</td><td></td><td>Tubing for water flow for zebrafish preparation</td></tr><tr><td>VaporGuard</td><td>VetEquip</td><td>931401</td><td>For recycling isoflurane</td></tr><tr><td>Vetbond tissue adhesive</td><td>3M</td><td>1469SB</td><td>To glue the glass window on the mouse skull, and to glue the laboratory tissue when preparing the fish.</td></tr><tr><td>XLPLN25XWMP2</td><td>Olympus</td><td></td><td>Multiphoton Excitation Dedicated Objective</td></tr></tbody></table>

deep-tissue three-photon fluorescence microscopy in intact mouse and zebrafish brain

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo imaging with subcellular resolution. 3PM has two major advantages for deep-tissue imaging over 2PM that has been widely used in biology laboratories: (i) longer attenuation length in scattering tissues by employing ~1,300 nm or ~1,700 nm excitation laser; (ii) less background fluorescence generation due to higher-order nonlinear excitation. As a result, 3PM allows high-contrast structural and functional imaging deep within scattering tissues such as intact mouse brain from the cortical layers to the hippocampus and the entire forebrain of adult zebrafish.
Today, laser sources suitable for 3PM are commercially available, enabling the conversion of an existing two-photon (2P) imaging system to a three-photon (3P) system. Additionally, multiple commercial 3P microscopes are available, which makes this technique readily available to biology research laboratories. This paper shows the optimization of a typical 3PM setup, particularly targeting biology groups that already have a 2P setup, and demonstrates intravital 3D imaging in intact mouse and adult zebrafish brains. This protocol covers the full experimental procedure of 3P imaging, including microscope alignment, prechirping of ~1,300 and ~1,700 nm laser pulses, animal preparation, and intravital 3P fluorescence imaging deep in adult zebrafish and mouse brains.

The successful completion of this protocol will result in a properly aligned microscope with optimal light parameters (e.g., pulse duration, NA) and animal preparations appropriate for in vivo 3PM. The commercially available 3P setup comprises appropriate mirrors and lenses for both ~1,300 nm and ~1,700 nm; therefore, no change in optics is required when the excitation wavelength is switched between 1,300 nm and 1,700 nm. If the lenses in a 3P setup do not have an appropriate coating for 1,300 and 1,700 nm, these need to be replaced with appropriate ones to reduce laser power loss. With the optimized 3PM and proper animal preparation, in vivo fluorescence and THG images with high contrast can be collected deep within the brains.
Figure 3&#160;shows representative 3P images of intact adult zebrafish. High-resolution, non-invasive, and deep imaging of genetically labeled neurons in the adult zebrafish brain is achieved using 3PM. Although imaging in the telencephalon region has been reported in the adult zebrafish brain using 2PM24,25,26,27, 3PM enables access to the entire telencephalon and regions that are more challenging or impossible to observe using other techniques. The distribution of cell layers in the optic tectum and cerebellum can be observed in Figure 3C. In a successful imaging session, the bone is visible in the THG channel and neurons visible in the fluorescence channel. For adult zebrafish imaging, the camera feature of the microscope was used to locate the fish (Figure 3A). This step is not necessary for mouse brain imaging as the glass window is large enough to make the brain easily accessible. High-resolution structural images of adult zebrafish brain were obtained using the system described in the previous sections. The skull is seen in the THG channel (Figure 3B), which helps navigate the brain and find the top surface. As observed in Figure 3C, neurons are distinguishable with a high signal-to-background ratio (SBR) deep in the adult brain. The tissue above the brain is visible in the fluorescence channel due to autofluorescence.
Figure 4&#160;shows multicolor 3P images of GCaMP6s-labeled neurons (green) and Texas Red-labeled blood vessels (red) together with THG (blue) signals in the adult mouse brain with 1,340 nm excitation10. The images are reproduced from previous work10. In Figure 4, the pulse energy at focus was maintained at ~1.5 nJ in the entire depth to obtain sufficient fluorescence and THG signals, and the maximum average laser power was ~70 mW. The pulse duration was adjusted to ~60 fs, and the effective NA was ~0.8. With optimization of the 3PM setup, high-contrast images were successfully obtained down to 1.2 mm from the brain surface, in the CA1 hippocampus region (Figure 4A,B). Figure 4C,D show Ca2+ activity traces of GCaMP6s-labeled neurons at a depth of 750 &#181;m for a 10 min recording session, showing high recording fidelity.
If the excitation laser is misaligned, nonuniformity in signal brightness across the field of view may be observed. Additionally, if the laser parameters, such as the pulse duration, the excitation pulse energy at focus, and the effective NA, are not optimized, the THG image from the fish skull or the craniotomy window of the mouse brain will not be visible and/or requires high excitation pulse energy (e.g., &#62;2 nJ/pulse at focus). Hence, the THG signals at the brain surface can be used as an indicator for an optimized 3PM setup before starting deep-tissue imaging.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63213/63213fig1v5.jpg" /> 
Figure 1: Schematic illustration of a 3PM setup. The wavelength of the excitation laser is set at ~1,300 nm or ~1,700 nm, output from the idler port of the NOPA. The prism-pair compressor and the Si plate compressor are used for the ~1,300 nm and ~1,700 nm laser, respectively, to prechirp the excitation laser pulse. The ~1,300 nm and ~1,700 nm laser beams can be switched with flipper mirrors. The signal port of the NOPA is used to obtain the triggering signal. A half waveplate and a PBS are used to control the excitation power. The fluorescence and THG are detected by GaAsP PMTs. Appropriate combinations of dichroic mirrors and bandpass filters are used to separate the fluorescence and THG signals. Abbreviations: 3PM = three-photon microscopy; NOPA = non-collinear optical parametric amplifier; PBS = polarizing beam splitter; THG = third-harmonic generation; DAQ = data acquisition; PMT = photomultiplier tube. <a href="https://www.jove.com/files/ftp_upload/63213/63213fig1v5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63213/63213fig2v5.jpg" /> 
Figure 2: Representative screenshots for intravital Imaging in fish (protocol section 4.1). (A) Camera mode view of the image acquisition software with 4x objective lens in place. Key features of the software are outlined and numbered as follows: 1. Imaging mode of the image acquisition software. The mode options are Camera, Multiphoton, and Multiphoton GG. For white light imaging with CCD camera, the Camera mode is chosen. 2. Clicking the Live button turns on the camera (or PMTs if in multiphoton options), and a real-time view of the microscope can be observed. 3. In Capture Setup tab, the desired imaging parameters (power, location, depth) are set. 4. In Capture tab, a folder location is assigned for the images to be saved. The imaging can be started in this tab. 5. The Z Control setting controls the depth of imaging by moving the z stage motor. 6. Representative image of a zebrafish head. The rostral side of the head is on the left. (B) Representative view of Camera mode with 25x objective lens. (C) Representative view of Multiphoton GG mode containing the THG image of image seen in (B). Abbreviations: CCD = charge-coupled device; PMT = photomultiplier tube; THG = third-harmonic generation. <a href="https://www.jove.com/files/ftp_upload/63213/63213fig2v5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63213/63213fig3v5.jpg" /> 
Figure 3: Representative images of the adult zebrafish brain acquired with the image acquisition software. (A) Camera mode image of adult zebrafish head acquired with a 4x objective lens. The top of the image is the rostral direction. The OT lobes and CB are outlined. (B) Representative image acquired in Multiphoton GG mode with a 25x objective lens containing the THG image of (A). (C) Fluorescence images of adult zebrafish brain at the intersection of the cerebellum and the optic tectum where GFP is expressed in cytoplasm of neurons in various depths. Scale bars = 100 &#181;m. Abbreviations: OT = optic tectam; CB = cerebellum; THG = third-harmonic generation; GFP = green fluorescent protein. <a href="https://www.jove.com/files/ftp_upload/63213/63213fig3v5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63213/63213fig4v5.jpg" /> 
Figure 4: Multicolor 3PM of GCaMP6s-labeled neurons (green), Texas Red-labeled blood vessels (red), and third-harmonic generation (blue) upon 1,340 nm excitation in the mouse brain. (A) Z-stack images down to 1,200 &#181;m from the brain surface with a field-of-view of 270 x 270 &#181;m (512 x 512 pixels per frame). The laser power was varied according to the imaging depth to maintain ~1.5 nJ pulse energy at the focus. Maximum average power under the objective was 70 mW. (B) Selected 2D images at various imaging depths. (C) Activity recording site at 750 &#181;m beneath the dura with a field-of-view of 270 x 270 &#181;m (256 x 256 pixels). (D) Spontaneous brain activity traces recorded in an awake mouse from the labeled neurons indicated in (C). The frame rate was 8.3 Hz, with a pixel dwell time of 0.51 &#181;s. The laser repetition rate was 2 MHz, and the average power under the objective lens was 56 mW. Each trace was normalized to its baseline and low-pass filtered using a hamming window of 0.72 s time constant. Scale bars = 50 &#181;m. This figure and the figure legend are reproduced from 10. Abbreviation: 3PM = three-photon microscopy. <a href="https://www.jove.com/files/ftp_upload/63213/63213fig4v5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63213/63213fig05.jpg" /> 
Figure 5: Effective attenuation length in the neocortex of the mouse brain. EAL (magenta line) is calculated from the tissue scattering (red line) and the water absorption in the tissue (blue line), assuming 75% water composition. The black stars indicate reported experimental data of EAL in the neocortex of the mouse brain3,21,28,29. Note that EAL varies in different tissues. Abbreviation: EAL = effective attenuation length. <a href="https://www.jove.com/files/ftp_upload/63213/63213fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Excitation wavelength 
			(nm)</td>
			<td>Immersion water</td>
			<td>Maximum laser power 
			(mW)</td>
			<td>Maximum pulse energy at focus* 
			(nJ)</td>
			<td>Typical EAL in the mouse cortex 
			(&#181;m)</td>
			<td>Imaging depth in the mouse cortex** 
			(mm)</td>
			<td>Pulse energy under the objective*** 
			(nJ)</td>
			<td>Maximum laser repetition rate**** 
			(MHz)</td>
		</tr>
		<tr>
			<td rowspan="4">1300</td>
			<td rowspan="4">H2O or D2O</td>
			<td rowspan="4"><img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />100</td>
			<td rowspan="4"><img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />2</td>
			<td rowspan="4">~300</td>
			<td>0.8</td>
			<td>~14</td>
			<td>~7</td>
		</tr>
		<tr>
			<td>1.2</td>
			<td>~55</td>
			<td>~2</td>
		</tr>
		<tr>
			<td>1.6</td>
			<td>~210</td>
			<td>~0.5</td>
		</tr>
		<tr>
			<td>2.1</td>
			<td>~1100</td>
			<td>~0.1</td>
		</tr>
		<tr>
			<td rowspan="4">1700</td>
			<td rowspan="4">D2O</td>
			<td rowspan="4"><img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />50</td>
			<td rowspan="4"><img alt="Equation 3" src="/files/ftp_upload/63213/63213eq03v2.jpg" />3</td>
			<td rowspan="4">~400</td>
			<td>0.8</td>
			<td>~7</td>
			<td>~7</td>
		</tr>
		<tr>
			<td>1.2</td>
			<td>~20</td>
			<td>~2.5</td>
		</tr>
		<tr>
			<td>1.6</td>
			<td>~55</td>
			<td>~1</td>
		</tr>
		<tr>
			<td>2.1</td>
			<td>~190</td>
			<td>~0.3</td>
		</tr>
	</tbody>
</table>
Table 1: Typical 3P excitation conditions for mouse cortex imaging. 
*With a high NA (~1.0) objective, pulse width of ~50 fs, and typical fluorophores such as fluorescent proteins (e.g., GFP and RFP). 
** With the assumption that the EAL is uniform in the entire cortex. 
*** To achieve ~1 nJ/pulse at focus, calculated from the EAL and the imaging depth. 
**** Calculated from the pulse energy under the objective and the maximum permissive laser power. 
Abbreviations: 3P = three-photon; NA = numerical aperture; GFP = green fluorescent protein; RFP = red fluorescent protein; EAL = effective attenuation length.

Watch this Scientific Journal Video about Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain at JoVE.com

Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain

Three-photon microscopy enables high-contrast fluorescence imaging deep in living biological tissues, such as mouse and zebrafish brains, with high spatiotemporal resolution.

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo ...

deep-tissue-three-photon-fluorescence-microscopy-intact-mouse

Research

JoVE Journal

Neuroscience

4.7K Views.  Cornell University. Three-photon microscopy enables high-contrast fluorescence imaging deep in living biological tissues, such as mouse and zebrafish brains, with high spatiotemporal resolution.

Video: Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain

Large-scale Scanning Transmission Electron Microscopy (Nanotomy) of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously applied large scale EM to human skin pancreatic islets, tissue culture and whole zebrafish larvae1-7. Here we describe a universally applicable method for tissue-scale scanning EM for unbiased detection of sub-cellular and molecular features. Nanotomy was applied to investigate the healthy and a neurodegenerative zebrafish brain. Our method is based on standardized EM sample preparation protocols: Fixation with glutaraldehyde and osmium, followed by epoxy-resin embedding, ultrathin sectioning and mounting of ultrathin-sections on one-hole grids, followed by post staining with uranyl and lead. Large-scale 2D EM mosaic images are acquired using a scanning EM connected to an external large area scan generator using scanning transmission EM (STEM). Large scale EM images are typically ~ 5 - 50 G pixels in size, and best viewed using zoomable HTML files, which can be opened in any web browser, similar to online geographical HTML maps. This method can be applied to (human) tissue, cross sections of whole animals as well as tissue culture1-5. Here, zebrafish brains were analyzed in a non-invasive neuronal ablation model. We visualize within a single dataset tissue, cellular and subcellular changes which can be quantified in various cell types including neurons and microglia, the brain&#39;s macrophages. In addition, nanotomy facilitates the correlation of EM with light microscopy (CLEM)8 on the same tissue, as large surface areas previously imaged using fluorescent microscopy, can subsequently be subjected to large area EM, resulting in the nano-anatomy (nanotomy) of tissues. In all, nanotomy allows unbiased detection of features at EM level in a tissue-wide quantifiable manner.

Large-scale Scanning Transmission Electron Microscopy Nanotomy of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution EM. Here we describe a universal method for nanotomy applied to investigate the zebrafish larval brain in health and upon non-invasive brain injury.

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously ...

Developmental Biology

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Biology

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Intravital Imaging of Fluorescent Protein Expression in Mice with a Closed-Skull Traumatic Brain Injury and Cranial Window Using a Two-Photon Microscope

The goal of this protocol is to demonstrate how to longitudinally visualize the expression and localization of a protein of interest within specific cell types of an animal's brain, upon exposure to exogenous stimuli. Here, the administration of a closed-skull traumatic brain injury (TBI) and simultaneous implantation of a cranial window for subsequent longitudinal intravital imaging in mice is shown. Mice are intracranially injected with an adeno-associated virus (AAV) expressing enhanced green fluorescent protein (EGFP) under a neuronal specific promoter. After 2 to 4 weeks, the mice are subjected to a repetitive TBI using a weight drop device over the AAV injection location. Within the same surgical session, the mice are implanted with a metal headpost and then a glass cranial window over the TBI impacting site. The expression and cellular localization of EGFP is examined using a two-photon microscope in the same brain region exposed to trauma over the course of months.

This study demonstrates delivery of a repetitive traumatic brain injury to mice and simultaneous implantation of a cranial window for subsequent intravital imaging of a neuron-expressed EGFP using two-photon microscopy.

The goal of this protocol is to demonstrate how to longitudinally visualize the expression and localization of a protein of interest within specific cell ...

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

Behavior Tracking and Neural Activity Pattern in the Zebrafish Forebrain

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used for measuring activities across brain regions through optical imaging methods. However, reports on the recording of brain activity in behaving adult zebrafish have been scarce. The present study describes procedures to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish. We focus on steps to restrain adult zebrafish from moving their heads, which provides stability that enables laser scanning imaging of the brain activity. The head-restrained animals can freely move their body parts and breathe without aids. The procedure aims to shorten the time of head restraint surgery, minimize brain motion, and maximize the number of neurons recorded. A setup for presenting an immersive visual environment during calcium imaging is also described here, which can be used to study neural correlates underlying visually triggered behaviors.

Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used ...