Two-Photon In Vivo Imaging of the Mouse Retina

Abstract

Source: Wang, Z., et al. Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina. J. Vis. Exp. (2021).

This video demonstrates a technique for in vivo retinal imaging using two-photon microscopy. Anesthetized mice are prepared for imaging by securing their heads, Applying eye lubricant, and placing a coverslip over the eye. Retinal ganglion cells (RGCs) expressing fluorescent proteins are first visualized using epifluorescence for alignment. Finally, two-photon imaging is employed, allowing for high-resolution visualization of RGCs.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation of mouse for image acquisition

1. Anesthetizing mouse for imaging
   NOTE: To mitigate exposure to isoflurane, ensure proper waste gas scavenging. Make sure the exhaust port of the anesthesia induction chamber is connected to a passive scavenging gas filter canister and the outlet of the mouse anesthesia nosepiece is connected to an active scavenging gas filter canister with a vacuum provided by a gas evacuation apparatus. Follow the manufacturer's guidelines for monitoring the filter canister weight for replacement.
   1. Anesthetize the mouse using the ketamine/xylazine cocktail. Alternatively, induce anesthesia via isoflurane inhalation if also using isoflurane for maintenance of anesthesia (imaging session > 30 min). Set the small animal anesthesia device to fill the induction chamber with 5% isoflurane mixed with room air at a flow rate of 0.5 L/min. Place the mouse in the induction chamber, and allow 15 s for the mouse to become anesthetized.
   2. Switch the isoflurane vaporizer to 0% and direct the anesthesia flow to the nosepiece. Perform a 5-second "O₂ flush" of the induction chamber. Take the mouse out of the induction chamber, and secure it in the head holder, attaching the nosepiece.
   3. Use a mixture of 1% isoflurane with room air at a flow rate of 0.5 L/min for maintenance of anesthesia. Monitor respiratory rate at 5-minute intervals throughout anesthesia, adjusting the isoflurane percentage to maintain a respiratory rate of ~60 breaths/min.
      ​NOTE: These settings (% isoflurane, flow rate) can also be used for the maintenance of anesthesia induced by the ketamine/xylazine cocktail.
2. Pupil dilation
   1. Prepare a solution of 1% w/v atropine and 2.5% w/v phenylephrine hydrochloride in water. Store the dilator solution at room temperature protected from light.
   2. Using a disposable pipette eye dropper, apply a drop of the dilator solution to each eye that will be imaged. Turn off the room lights, and wait 5 min for the pupil to dilate. When the pupil is dilated, blot away the dilator solution with lint-free tissue.
      NOTE: Ensure dilator solution does not enter the nostrils.
   3. Apply a large drop of lubricant eye gel to each eye that will be imaged. If both eyes are to be imaged, apply a small piece of plastic cling film over the eye gel on the non-imaging eye to prevent dehydration. If only imaging one eye, apply lubricant eye ointment to the eye that will not be imaged.
3. Positioning mouse for imaging
   1. To secure the mouse in the imaging head holder, rotate the main arm of the head holder until the earpiece bar is tilted at an angle of 60° or more below horizontal. Secure the lower ear canal pin in the inward extended position and the upper ear canal pin in the withdrawn position.
   2. With the mouse facing the bite bar, mount one ear onto the extended lower pin, inserting the pin into the ear canal. Loosen the screw securing the upper ear canal pin, and extend the pin into the other ear canal. Tighten the screw to secure the head.
   3. Slide the bite bar toward the head of the mouse. Gently elevate the head of the mouse, then lower the maxillary incisors of the mouse into the hole of the bite bar. Retract the bite bar with gentle force to secure the head of the mouse, and secure the bite bar position by tightening the screw.
   4. If using isoflurane, slide the nosepiece through its slot onto the bite bar prior to securing the incisors of the mouse. Secure and tighten the bite bar position as in step 1.3.3. Tighten the nosepiece using the two screws on its upper face until it fits snugly on the nose of the mouse, but does not constrict the snout.
   5. Transfer the mouse, in the head holder, to the microscope stage beneath the objective. Rotate the main arm of the head holder until the pupil of the eye is oriented straight up, in line with the light path (Figure 1).
   6. Place a #1.5 coverslip in the compact filter holder and attach the holder to the microscope stage. Lower the coverslip toward the eye, contacting the lubricant eye gel, so that it lies horizontally immediately above the cornea (Figure 1). Ensure that the coverslip does not touch the cornea.
      NOTE: On the microscope, ensure the excitation light path is set to epifluorescence and the emission light path is set to the eyepiece. Turn on the epifluorescence illuminator at its lowest power and open the illuminator shutter. Choose the epifluorescence illuminator wavelength and fluorescence filter corresponding to the fluorophore being imaged.
   7. Maneuver the stage in the X-Y dimensions and objective in the Z-position using the stage control and motorized focus drive until the widefield excitation light fully covers the cornea. Looking through the eyepiece, continue to adjust the Z-position of the stage until the fluorescent cells or structures in the retina come into focus. Increase epifluorescence illuminator power if the sample signal is insufficiently bright to resolve individual cells or structures of interest through the eyepiece.
      NOTE: If having trouble locating the retina, the iris is a high-contrast landmark to anchor on and then focus down toward the retina. This step will also allow verification that the pupil is maximally dilated.
   8. Obtain an imaging area on the axis with the mouse lens. Adjust the angle of the mouse using the various degrees of freedom on the head holder until only expansion or contraction of out-of-focus light occurs when adjusting the focal plane. Turn off the epifluorescence illuminator and close the illuminator shutter.
      NOTE: Significant X-Y parallax of out-of-focus light while scrolling through the Z-direction focal planes indicates that the retina is not on the axis with respect to the imaging light path.

2. Two-photon image acquisition

1. Image setup and acquisition parameters
   ​NOTE: Turn off room lights, and cover stray light sources in the room. Ensure the epifluorescence illuminator is off, and the illuminator shutter is closed.
   1. Switch the excitation light path to the laser and the emission light path to the photomultiplier tubes (PMTs).
   2. In the image acquisition software, set a frame size of 512 x 512 and a frame average of 3. Set the steps per slice to -8 µm. Designate z-stepping to start at the top of the stack and progress downward, minimizing two-photon laser activation of photoreceptors.
      ​NOTE: The step size can be reduced to increase Z-resolution at a cost of increased imaging time, but 8 µm steps are sufficient to resolve cell somata in this imaging configuration.
   3. Turn on and enable the PMTs. Adjust the PMT voltage to 680 V. Enable the excitation shutter.
      ​NOTE: Powered PMTs are susceptible to light damage. Ensure that the epifluorescence illuminator is off, the microscope enclosure curtain is drawn, and the room lights are off.
   4. Begin a live image preview of the target tissue, starting with 1% laser power. Auto-adjust the display brightness to visualize the cells or structures of interest. Auto-adjust the scan phase. If the target tissue is dim or unclear, increase the laser power percentage until structures become visible without surpassing the limit set in step 2.4.4, which corresponds to 45 mW.
      ​NOTE: For a 16-bit image, a display value of ~1000 when auto-adjusting brightness in a Z-plane containing structures of interest indicates sufficient brightness of the sample.
   5. Maneuver the microscope stage in the X-Y direction to center on a desired imaging area, then navigate to the Z-plane with the structures of interest in focus.
      ​NOTE: Imaging adjacent to the optic nerve head allows it to serve as an unambiguous landmark for chronic imaging experiments.
   6. If this is a chronic time-lapse experiment, open a previous image on the acquisition computer and use it as a reference to find the same area of interest. Ensure that the angle of imaging is similar to that of previous images to acquire the same set of cells with minimal parallax.
      NOTE: To image the same cells as in previous time points (Figure 2), further adjustment of the mouse head position is likely required.
   7. Set the Z-limits of the imaging stack by navigating to the uppermost and lowermost Z-planes of interest, and acquire the image. Upon completion of image acquisition, disable the PMTs and the emission shutter. Switch the emission light path back to the eyepiece, and the excitation light path to epifluorescence illumination. Remove the mouse from the microscope stage.

Results

Figure 1: Positioning mice for in vivo imaging. To position the mouse with the pupil on the axis with the light path, the anesthetized mouse is first restrained in a head holder, the head is rotated and angled...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
#1.5 coverslip	ThermoFisher	152440	Richard-Allan #1.5 24 mm x 40 mm
50 mL glass syringe	Hamilton	80950	22G cemented needle
Anesthesia Air Pump	RWD Life Science	R510-30
Atropine	Sigma	A0132	For pupil dilator solution
Basic Small Animal Anesthesia Device	RWD Life Science	R500IE
Borosilicate glass capillary	Sutter	B150-86-10	Outside diameter 1.50 mm, inside diameter 0.86 mm, length 10 cm
CFP/YFP filter cube	Chroma	custom	480/40, 505 long pass, 535/30
ChromoFlex - Two channel PMT detection unit	Scientifica	S-MPLG-1002
Circulating heating pump	Braintree Scientific	tp-700	Set to 37 °C
Cling film	VWR	10713-916
Compact Filter Holder	ThorLabs	DH1	Holds coverslip over mouse eye
Cx3cr1-GFP transgenic mice (B6.129P2(Cg)-Cx3cr1tm1Litt/J)	The Jackson Laboratory	5582
DAQ controller chassis	National Instruments	PXIe-1073
Data acquisition device	National Instruments	BNC-2090A
FPGA module with digitizer	National Instruments	NI-5734
Gas Evacuation Apparatus	RWD Life Science	R546W
GenTeal Severe lubricant eye gel	Alcon	(from local pharmacy)	For use during imaging
GFP/Red filter cube	Chroma	custom	535/30, 560 long pass, 605/70
Heating pad	McKesson Medical and Surgical	190147
HyperScope Launch Optics for use with Pockels Cell	Scientifica	S-MP-101080
HyperScope Main module	Scientifica	S-MP-100466
HyperScope Scan Path	Scientifica	S-MP-100406
HyperScope X galvo Module	Scientifica	MP-100443
ImageJ Fiji software		Freeware
Isoflurane	Patterson Veterinary	NDC 14043-704-06
Isoflurane gas filter cannister (active scavenging)	RWD Life Science	R510-31
Isoflurane gas filter cannister (passive scavenging)	RWD Life Science	R510-31S
ketamine HCl (100 mg/mL)	Vedco	NDC 50989-161-06
M32 to M26 adapter	ThorLabs	M32M26S
MaiTai GUI software	Spectra-Physics	NA
MATLAB software	MathWorks	NA	R2015b
meloxicam (5 mg/mL)	Boehringer Ingelheim	NDC 0010-6013-01	Analgesic
Micorscope Objective	Edmund Optics	46-404	Mitutoyo WE715042319
micropipette puller	Sutter	Flaming/Brown Model P-97
Mineral oil	Fisher	BP2629-1
Mini bulldog hemostatic clamp	Fine Science Tools	18053-28
Miniature EVA Tubing 0.02" ID, 0.06" OD	McMaster Carr	1883T1
Miniature EVA Tubing 0.05" ID, 0.09" OD	McMaster Carr	1883T4
Mouse head holder	Narishige	SGM-4
No. 5 Forceps	Fine Science Tools	11251-10
Optic Posts 1/2"	ThorLabs	TR3-P5
Optical power meter kit	ThorLabs	PM100D
pE-300 Ultra LLG Deivery	Scientifica	COO-LED3ULLGs
Phenylephrine hydrochloride	Sigma	P6126	For pupil dilator solution
Pockels cell	Conoptics	350-80-02
Pockels cell amplifier	Conoptics	Model 302RM
Proparacaine hydrochloride	Sigma	1571001	For eye immobilization
Red & Far Red short pass filter Cube	Chroma	custom	560 short pass
Rotating 1/2" post clamp	ThorLabs	SWC
ScanImage package	Vidrio Technologies	Freeware	Image acquisition software; Version 5.4.0 (2018); requires MATLAB
sodium chloride solution, sterile (0.9%)	Fresenius Kabi	NDC 63323-186-01
Stereomicroscope	Leica	S9 E
Tabletop centrifuge	Oxford	Benchmate C8
Terramycin oxytetracycline/polymyxin B antibiotic ophthalmic ointment	Zoetisus	NA	For use after intravitreal injection
ThermoRack cooling system	Solid State Cooling Systems	ThermoRack 401	Set to 20 °C
Ultrafast Ti:Sapphire laser	Spectra-Physics	Mai Tai DeepSee
Vgat-Cre transgenic mice (Slc32a1tm2(cre)Lowl/J)	The Jackson Laboratory	16962
VGlut2-Cre transgenic mice (Slc17a6tm2(cre)Lowl/J)	The Jackson Laboratory	16963
VivoScope for In Vivo Imaging	Scientifica	S-MPVS-1200-00P
White petrolatum-mineral oil lubricant eye ointment	Stye	NA	For use after imaging
xylazine HCl (20 mg/mL)	Akorn	NDC 59399-110-20