This method describes a chronic preparation that allows optical access to the hippocampus of living mice. This preparation can be used to perform longitudinal optical imaging of neuronal structural plasticity and activity-evoked cellular plasticity over a period of several weeks.
Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from subcellular to network levels and at temporal scales from milliseconds to weeks. In addition, two-photon imaging can be combined with a variety of behavioral tasks to explore the causal relationships between brain function and behavior. However, in mammals, limited penetration and scattering of light have limited two-photon intravital imaging mostly to superficial brain regions, thus precluding longitudinal investigation of deep-brain areas such as the hippocampus. The hippocampus is involved in spatial navigation and episodic memory and is a long-standing model used to study cellular as well as cognitive processes important for learning and recall, both in health and disease. Here, a preparation that enables chronic optical access to the dorsal hippocampus in living mice is detailed. This preparation can be combined with two-photon optical imaging at cellular and subcellular resolution in head fixed, anesthetized live mice over several weeks. These techniques enable repeated imaging of neuronal structure or activity-evoked plasticity in tens to hundreds of neurons in the dorsal hippocampal CA1. Furthermore, this chronic preparation can be used in combination with other techniques such as micro-endoscopy, head-mounted wide field microscopy or three-photon microscopy, thus greatly expanding the toolbox to study cellular and network processes involved in learning and memory.
In mammals, the hippocampus is a key brain region for the encoding and recall of episodic memories as well as for spatial navigation1,2,3,4. For this reason, the hippocampus has been – and still is – a very important model to study the basic mechanisms that allow the brain to encode and recall memories5,6,7 or to navigate in an environment8,9 collecting rewards and avoiding dangers. In addition, the hippocampal formation is one of the brain regions where new neurons are generated throughout the life of rodents10,11 and, possibly, of humans12,13. Finally, degeneration or impairment of the hippocampal formation are associated with neurological and psychiatric disorders, including Alzheimer’s disease14.
In mice, the hippocampus is located approximately 1 mm below the brain surface15. Its position has prevented optic access in the intact brain and consequently, longitudinal studies of hippocampal dynamics have relied mostly on magnetic resonance (MR) imaging, electrophysiology, and ex vivo imaging analyses. MR imaging methods allow tracking of biological processes (e.g., gene expression changes16) in the same animal over multiple days, but lack the spatial resolution to discriminate single neurons. Classic in vivo electrophysiological techniques offer very high temporal resolution and exquisite sensitivity to changes in membrane potential. However, they have a limited spatial resolution and they lack the ability to reliably track the same cells over longer time periods. Optical imaging allows more diverse processes to be studied by virtue of its high temporal and spatial resolutions. However, ex vivo imaging only provides snapshots of ongoing processes, and thus it is not suitable for longitudinal studies during which the animals learn and recall information.
In vivo optical imaging combines some advantages of MR imaging and electrophysiology with those of optical imaging. Therefore, it is very well suited for longitudinal and correlative analyses of mouse brain dynamics and behavior. This is relevant in studies of biological processes with very fast (milliseconds to seconds) or very slow (days to weeks) time scales. Examples for such processes that are relevant for neuroscience are membrane voltage dynamics, Ca2+ transients, cellular plasticity and structural changes, which are all believed to be very important for memory formation and recall. Different methods have extended in vivo imaging to the dorsal hippocampus18,19,20,21,22. Acute preparations have allowed the tracking of pyramidal neuron (PN) activity as well as their dendrites and dendritic spines for several hours20,22. This temporal timeframe, however, does not allow long-term structural changes, which might underlie incremental learning, to be studied. Chronic preparations – in combination with micro-endoscopes23,24 or with long working distance (WD) standard microscope objectives21 – have enabled repeated imaging of the dorsal hippocampus over several weeks.
Here, we describe a chronic preparation that provides recurrent optic access to the CA1 sub-field of the dorsal hippocampus of living mice using a permanently inserted imaging cannula. This preparation allows repeated access to the CA1 without functional disturbance and is suitable for intravital two-photon (2P) or wide-field epifluorescence imaging. Two examples of 2P deep brain chronic imaging in the dorsal CA1 of live mice are detailed: longitudinal imaging of dendritic structure and dendritic spine dynamics and longitudinal imaging of activity-evoked plasticity. The salient advantages and limitations of the technique are discussed.
All of the methods described have been approved by the Government of Upper Bavaria (licence 2016_ROB-55.2Vet-2532.Vet_02-16-48) and by the Stanford and Max Planck Florida Institute for Neuroscience Administrative Panels on Laboratory Animal Care.
1. Preparation of the imaging cannula
2. Implantation of the imaging cannula over the dorsal hippocampus
3. Postoperative care
4. Preparation of the imaging session
5. Imaging session
Since the cannula is placed just dorsal to CA1, the dorsal aspect of the CA1 is more proximal to the microscope if compared to the ventral one. The alveus is the most proximal structure followed in order by stratum oriens (SO), stratum pyramidale (SP), stratum radiatum (SR) and stratum lacunosum moleculare (SLM), the most distal layer (Figures 1B, C). Longitudinal 2P imaging of neuronal structure with subcellular resolution and of activity-evoked plasticity with cellular resolution are presented as representative results. To excite the fluorophores green fluorescent protein (GFP), d2Venus and TurboFP635, a femtosecond pulsed Ti:Sapphire laser tuned to 920 nm is used and the average laser power is adjusted to 5-25 mW at the sample. The different emission wavelengths are separated using emission filters and different photomultiplier tubes.
Longitudinal imaging of dendritic structure and dendritic spines dynamics.
To sparsely label hippocampal PNs and visualize their structure, a Thy1-GFP transgenic mouse line (M line) is used. Thy1-GFP mice express cytoplasmic enhanced GFP under the control of the Thy1 promoter in a sparse, random population of PNs25. Generally, the major axis of CA1 PNs is roughly perpendicular to the XY-imaging plane (Figures 1B, Figure 2A and Supplementary Movie 1). Basal dendrites extend in the SO, from the soma towards the cannula, whereas apical and apical tuft dendrites extend distally to the cannula, in the opposite direction (Supplementary Movie 1). Since the preparation leaves the deepest fibers of the alveus intact, a few fluorescent fibers traversing the field of view, just beneath the cannula, should be visible (Figure 2A and Movie 1). The preparation allows imaging of PN dendrites with spine resolution (Figure 2 A-C). To image dendrites and dendritic spines, a 25X 1.0 NA, 4 mm WD, water immersion commercial objective lens is used.
For longitudinal tracking, several brain regions within the field of view of the cannula are defined during the first imaging session. Each region corresponds to an area of approximately 240 x 240 µm and contains between 1 and 7 dendritic segments (Figure 2B). These regions are manually mapped to a low magnification three-dimensional stack showing the pattern of GFP expression in the volume below the imaging cannula (Figure 2A). Then, 1 µm z-step image stacks of CA1 PN basal dendrites are acquired at different time intervals (from 24 h to 3 days) for up to about 14 days (Figure 2C). Longer imaging durations and intervals are possible26. Each imaging session lasts approximately 60 to 90 min. Although most images are of dendritic spines in the SO, it is also possible to image dendritic spines in the oblique dendrites of SR (Figures 2D-F). In addition to spine density, this method enables the study of spine dynamics by quantifying their survival, gain and loss rates26,27,28,29,30,31,32. To score and track dendritic spines over time (Figure 2C), a custom MATLAB interface is used. This enables the alignment of the image stacks acquired at different time points and supports manual labeling of dendrites and spines while tracking dendritic lengths and spine positions over time26. Importantly, this method can be used to distinguish (per each time point, excluding the first one) between pre-existing and newborn dendritic spines. This is important as the different classes of dendritic spines are thought to have different roles in memory acquisition and retention33.
Longitudinal imaging of activity-evoked plasticity.
To image activity-evoked plasticity in CA1 PNs, the dorsal CA1 hippocampal area is injected with a viral vector expressing green fluorescent destabilized d2Venus via an enhanced form of the synaptic activity-responsive element (E-SARE) within the Arc enhancer/promoter and red fluorescent TurboFP635 via the ePGK promoter34. This allows for imaging levels of activity-evoked plasticity of hundreds of CA1 PNs in each animal35. Given the very dense labeling of PNs, it is generally not possible to resolve the dendrites of CA1 PNs (Supplementary Movie 2).
To image the somata of CA1 PNs, a 40X 0.8 NA, 3 mm WD, water immersion objective lens is used. For longitudinal tracking, 1 to 9 brain regions are defined per mouse during the first imaging session. Each region corresponds to an area of approximately 300 x 300 µm and contains between 50 and 150 cells (Figure 3A). These regions are manually mapped to local tissue landmarks visible at a lower magnification. Then, 3 µm z-step image stacks are acquired, which encompass the SP of CA1 PNs (Figure 3A and Supplementary Movie 2) at different time intervals (from 24 h to 6 days) for up to about 30 days. Each imaging session lasts approximately 60 to 90 min. E-SARE activation peaks 6 to 8 h after an exposure to a new or enriched environment (EE, Figure 3B) and decays over the course of a few days. Thus, we generally image 6 to 8 h after experience and allow for 5 days between imaging sessions35.
To quantify d2Venus and TurboFP635 fluorescence values, a circular region-of-interest 4.64 µm in diameter is drawn, which is smaller than a neural cell body, centered to the cell soma. We then progress to the next time point, score the soma of the same cell in the same way, and iterate this procedure for all time points and all visible cells in the longitudinal dataset. The mean value of each neuron’s (activity-dependent) d2Venus emission is normalized by its mean (activity-independent) TurboFP635 emission. This method enables the investigation of long-term dynamics of ensemble plasticity of CA1 PNs35 (Figure 3C).
Figure 1: Preparation for in vivo deep brain optical imaging. (A). Top (left) and side (right) views of example imaging cannuals. Imaging cannulas have a transparent glass bottom to allow optical access to the hippocampus. (B). Schematic description of the preparation highlighting the relative position of the imaging cannula, a CA1 PN and the three layers of fibers from dorsal to ventral. (C, D). Schematic description of the imaging setup (C) and of the animal fixation (D) during an imaging session. Please click here to view a larger version of this figure.
Figure 2: Longitudinal imaging of dendritic structure and dendritic spines dynamics. (A). 2P image stack (Maximum Intensity Projection (MIP) of 59 image planes, 2 µm z-spacing) of neurons and dendrites labelled by GFP in a live Thy1-GFP mouse. (B). Higher magnification (MIP of 53 image planes, 1 µm z-spacing) detailing basal dendrites located in SO. (C). Time-lapse image sequence of a dendritic segment imaged over 14 days. Arrowheads indicate dendritic spines tracked over 14 days. (D, E). 2P image stack of neurons and dendrites labelled by GFP in a live Thy1-GFP mouse; (D) ZY projection (31 image planes, 3 µm z-spacing) and (E) XY projections (17 image planes, 3 µm z-spacing). (F). Higher magnification (single image plane) detailing apical dendrites and dendritic spines located in SR. Arrowheads indicate dendritic spines. Excitation: 920 nm; emission peak: 510 nm. Scale bars: A, 50 µm; B, 10 µm; C, 2 µm; D and E, 15 µm; F, 4 µm. Please click here to view a larger version of this figure.
Figure 3: Longitudinal imaging of activity-evoked plasticity. (A). 2P images (single image planes) from a live mouse, showing the same cells on Baseline Day 0 and after EE on Day 1. (B). 2P image stacks (MIPs of 4-6 image planes, 3 µm z-spacing) showing E-SARE activation patterns specific for environment A (Days 1, 19 and 25) and environment B (Days 7 and 13). Green: d2Venus fluorescence. Red: TurboFP635 fluorescence. Excitation: 920 nm; d2Venus emission peak: 530 nm; TurboFP635 emission peak: 635nm. Scale bars: A, 20 µm; B, 10 µm. This figure has been modified from Attardo et al., 201835. Please click here to view a larger version of this figure.
Figure 4: Imaging of neuronal structure in hippocampal DG using three-photon (3P) microscopy. (A-H). 3P images (single image planes) of neurons and dendrites labelled by GFP in a live Thy1-GFP mouse detailing (A-E) PNs in the CA1 and (F-H) granule cells in the DG. Excitation: 1400 nm; emission peak: 510 nm. Scale bar: 40 µm. Please click here to view a larger version of this figure.
Supplementary Movie 1: Imaging field of view in a live Thy1-GFP mouse. 2P image stack (83 image planes, 7 µm z-spacing) of neurons and dendrites labelled by GFP (white) in a live Thy1-GFP mouse extending from the bottom of the cannula to SLM. To account for the decay of fluorescence signal with increasing depth, we used a non-linear gradient of photomultiplier tubes’ gain. Please click here to download this file.
Supplementary Movie 2: Imaging of activity-evoked plasticity. 2P image stack (28 image planes, 3 µm z-spacing) of neurons expressing E-SARE reporter of IEG expression in a live mouse encompassing SP. Green: d2Venus fluorescence. Red: TurboFP635 fluorescence. Please click here to download this file.
Here, a procedure for repeated 2P imaging of the dorsal CA1 in live mice is described. After the surgery, the mouse usually recovers within 2 days. The procedure induces minimal astrogliosis26,43. Hemorrhage and edema which might follow the surgery are usually re-adsorbed within 10 to 14 days. Generally, from 14 days post-implantation onwards the preparation is sufficiently clear to perform intravital imaging. The success of the surgery does not depend on working in a sterile environment. However, it is crucial to maintain a high level of hygiene, to avoid complications due to surgery-associated infections. This is obtained by meticulously cleaning the surgical instruments before and after the surgery and by heat-sterilizing them immediately prior to each usage (step 2.1.1). The optic cannula is kept into a clean, sterilized container and rinsed with sterile saline just before the implantation. Performing common surgical practices of hands disinfection and cleaning of the surgical station is also very important. The preparation remains stable and allows cellular and subcellular resolution imaging for several weeks26,35.
Critical steps, modifications and troubleshooting.
It is important to peel the external capsule until the deepest fibers are exposed. Failure to expose the alveus might result in the inability to focus on the soma of PNs, or in reduced resolution imaging dendritic spines, when using commercial objectives with 3- or 4-mm WD. To this aim, it is useful to ablate the neocortex very slowly using a 0.9 mm diameter needle and then switch to a 0.3-0.5 mm diameter (24-29 gauge) needle for a finer control of suction when removing the most dorsal fibers. Alternatively, fine forceps may be used to remove the remaining cortex after fiber exposure36.
Bleeding during surgery can be problematic, as blood obstructs the view. Waiting for the clot to form and then rinsing with saline to wash away residual blood is recommended. Repeat as necessary.
A snug fit between the cannula and the craniotomy helps increase the stability of the preparation by keeping the cannula in place before application of the cement, especially if the outer rim of the cannula is flush with the skull. Since the sizes of the trephine drill and the cannula are matched, a loose fit can arise because of irregularities on the side of the cannula – which require slightly larger craniotomies to fit (see step 2.3.14) – or from an irregular craniotomy. Any cannula irregularities must be filed off (steps 1.3 and 1.12) and the trephine must be held perpendicular to the skull until the craniotomy is completed (step 2.3.12). Removing the trephine from the skull before the craniotomy is completed may result in irregular craniotomies.
Limitations-Invasiveness and stability of the preparation.
It is difficult to evaluate the effect of cortical ablation as it is arduous to precisely define the areas affected directly and indirectly. In general, the surgery removes part of the parietal cortex and part of the visual and hindlimb sensory cortex21. The ablated cortex does not directly project to the hippocampus and hippocampal tissue is neither touched nor injured. Importantly, it has been shown that implantation of an imaging cannula does not grossly alter hippocampal function and specifically hippocampal-dependent learning21,36,37,38,39. Still, it would be important to quantify to what extent both the cannula and the external part of the implant (head holder plate and dental acrylic cap) are chronic stressors by assessing corticosterone blood levels and adrenal gland weight in comparison to unimplanted mice.
The preparation generally remains stable from weeks to months26. In the long term, skin and bone growth tend to displace the acrylic cap and to increase the instability of the imaging preparation.
Optical limitations.
Conventional 2P microscopy allows imaging up to about 1 mm deep into neocortical tissue40,41. Consistent with this, it is possible to image dendrites and dendritic spines located in the SR (Figure 2D-F) or SLM36. However, imaging through a cannula poses limitations to the effective NA. To achieve the maximum resolution, the diameter and depth of the imaging cannulas should be matched to the imaging NA, as smaller diameters and longer depths will clip light of high NA objectives. For instance, when imaging with a 1.0 NA water immersion objective through a 1.6 mm long cannula, a 3.65 mm inner diameter is needed to keep the full NA. However, using a cannula of this diameter will increase the compression on the hippocampus and might affect the health of the tissue, for this reason, we use a cannula with a smaller diameter. When imaging with a 0.8 NA water immersion objective through a 1.6 mm long cannula, an inner diameter of 2.5 mm would be sufficient to keep the full NA. However, 0.8 NA water immersion objectives have a shorter WD (3 mm in our case), which can prevent from focusing at the SP.
These calculations apply to the center of the field of view at the bottom of the cannula. However, moving the imaging field of view sidewise – closer to the edges of the cannula – or focusing deeper into the tissue – farther from the glass surface of the cannula – further decreases the effective NA at the focal plane and thus reduces resolution. This will lead to non-homogenous resolution across the different volumes of imaged tissue and can be a concern for quantitative imaging at subcellular resolution, especially when using super-resolution techniques such as 2P-STED microscopy42. These issues are less important when imaging at cellular resolution.
Tissue motion.
Motion within the tissue – originating from breathing and heartbeat in anesthetized animals – tends to become more severe with increased distance from the imaging cannula. This is possibly because the imaging cannula applies mechanical pressure to the brain thus counteracting some of the motion in the vicinity of the cannula (similarly to neocortical preparations). Thus, although imaging of dendritic spines is possible in SR and SLM, in our hands, it is most robust dorsal to the SO up to ≈200 µm from the surface of the cannula. To compensate for motion, we use resonant scanners and offline averaging. Several images (4 to 6 repetitions) are acquired per image plane of a z-stack at the maximum available speed (30 frames/s). All the repetitions for each z-plane are then deconvolved (using the commercial software, AutoQuant), registered (using ImageJ) and averaged into a single image26. For imaging of somata, motion is often negligible upon anesthesia35 and two averages are often sufficient to compensate for motion artifacts.
Future applications or directions of the method.
The preparation can be combined with micro-endoscopes26,43. Micro-endoscopes are rigid optic probes which use gradient refractive index (GRIN) microlenses to guide light to and from deep tissue18. The use of micro-endoscopes allows cannulas of smaller diameters or even no cannulas at all. However, commercial micro-endoscopes are less well corrected for optical aberrations and have lower NA than commercial objectives. Current probes reach lateral and axial resolutions of ≈0.6-1 µm, ≈10-12 µm, respectively17,18,44. The use of micro-endoscopes also enables the combination of this preparation with head-mounted integrated widefield microscopes45,46,47.
The method lends itself also to use in non-anesthetized mice, and it has been used to investigate cellular activity using Ca2+ sensors in awake head-fixed mice21,37,48,49. In these cases, due to the fast time scales of the fluorescence changes, it is advisable to implement line registration50. It is also possible to adapt the preparation for imaging of other hippocampal sub-regions such as the dentate gyrus (DG)39,51,52. Combining this preparation with 3P excitation53,54 with 1 MHz frequency pulsed laser tuned to 1400 nm, we were able to image deeper into the hippocampal formation reaching the molecular layer, granule cell layer and the hilus of the DG (Figure 4) without removing the overlaying CA1.
In conclusion, we present a method that provides optical access to the dorsal hippocampus and allows longitudinal and correlative studies of the dynamics of hippocampal structure and activity. This technique extends the possibilities of analysis of hippocampal function under physiological and pathological conditions.
The authors have nothing to disclose.
U. A. F. is supported by the Schram foundation; C.-W. T. P. and W. G. are supported by the Max Planck Society; L.Y. and R.Y. are supported by the Max Planck Society and National Institute of Health (R01MH080047, 1DP1NS096787); A. C. is supported by an FP7 Grant from the European Research Council, the ERANET and I-CORE programs, the Chief Scientist Office of the Israeli Ministry of Health, the Federal Ministry of Education and Research, Roberto and Renata Ruhman, Bruno and Simone Lich, the Nella and Leon Benoziyo Center for Neurological Diseases, the Henry Chanoch Krenter Institute for Biomedical Imaging and Genomics, The Israel Science Foundation the Perlman Family, Adelis, Marc Besen, Pratt and Irving I. Moskowitz foundations; A. A. is supported by the Max Planck Society, the Schram foundation and the Deutsche Forschungsgemeinschaft (DFG). The 3P images were acquired during the Advanced Course on Neuroimaging Techniques at the Max Planck Florida Institute for Neuroscience. The Advanced Course on Neuroimaging Techniques is supported by the Max Planck Society, the Florida State Max Planck Scientific Fellowship program and by the Max Planck Florida Institute Corporation Partnership program. We would like to thank Thorlabs, Coherent and SpectraPhysics for providing support and equipment for the 2P / 3P imaging system during the course. We are also grateful to Henry Haeberle and Melissa Eberle for assistance with the system during the course.
Professional drill/grinder IBS/E | Proxxon GmbH | 28481 | Pecision drill |
MICROMOT drill stand MB 200 | Proxxon GmbH | 28600 | Movable ruler table |
MICRO compound table KT 70 | Proxxon GmbH | 27100 | Movable ruler table |
Machine vice MS 4 | Proxxon GmbH | 28132 | Movable ruler table |
Stainless steel tube Ø 3,0 x 0,25 mm (Inner Ø 2,5 mm ) L = 500 mm | Sawade | R00303 | Stainless steel tube for the cannula metal ring |
Microscope Cover glass (4 mm round) | Engelbrecht Medizin and Labortechnik | Glass coverslips for the cannula glass | |
Schlusselfeilensatz 6-tgl. Im Blechetui | Hoffmann Group | 713750 160 | Manual files |
Präzisions-Nadelfeile Gesamtlänge 140 mm 4 | Hoffmann Group | 527230 4 | Manual files |
UV-Curing Optical Adhesives | Thorlabs | NOA81 | UV-curing adhesive |
UV Curing LED System, 365 nm | Thorlabs | CS2010 | UV-curing LED driver unit |
Stemi 305 | Zeiss | Stereoscope | |
Presto II | NSK-Nakanishi Germany | Z307015 | Dental drill |
Diamantbohrer FG (5 St.), Zylinder flach, 837-014 fein | MF Dental | F837.014.FG | Files for the dental drill |
Diamantbohrer FG (5 St.), Zylinder flach, 837-014 grob | MF Dental | G837.014.FG | Files for the dental drill |
Graefe Forceps – Straight / Serrated | Fine Science Tools | 11050-10 | Forceps for the surgery |
Burrs for Micro Drill | Fine Science Tools | 19008-05 | 0.5 mm width burr for the micro-drill |
Burrs for Micro Drill | Fine Science Tools | 19008-09 | 0.9 mm width burr for the micro-drill |
MicroMotor mit Handstück | DentaTec | MM11 | Micro-drill for the craniotomy |
Dumont #3 Forceps | Fine Science Tools | 11231-30 | Dumont forceps for the surgery |
Fine Scissors – ToughCut | Fine Science Tools | 14058-09 | Scissors for the surgery |
Trephine | MW Dental | 229-020 | Trephine drill – 3.0 mm diameter; for the micro-drill |
Stainless Steel Self-Tapping Bone Screws | Fine Science Tools | 19010-10 | 0.86 mm width bone screws |
Stereotaxic apparatus | Kopf | Stereotaxic apparatus | |
3-D-Gelenkarm | Hoffmann Group | 442114 | Stereotaxic arm and plate holder |
Aufnahme 2SM | Hoffmann Group | 442100 2SM | Stereotaxic arm and plate holder |
Hot Bead Sterilizers | Fine Science Tools | 18000-45 | Glass beads sterilizer |
Isofluran CP, Flasche 250 ml | Henry Schein VET GmbH | 798932 | Liquid isoflurane for anesthesia |
Harvard Apparatus Isoflurane Funnel-Fill Vaporizer | Harvard Apparatus GmbH | 34-1040 | Isoflurane vaporizer |
Lab Active Scavenger | Gropper Medizintechnik | UV17014 | Isoflurane scavenger system |
Metacam 0,5% Injektionslsg. (Hund / Katze), Flasche 20 ml | Henry Schein VET GmbH | 798566 | Meloxicam, anti-inflammatory |
Vetalgin 500 mg/ml | MSD Tiergesundheit | Vetalgin, pain killer | |
CMA 450 Temperature Controller | Hugo Sachs Elektronik – Harvard Apparatus GmbH | 8003770 | Heating blanket |
Bepanthen Augen- und Nasensalbe | Bayer AG | Ophtalmic ointment | |
KL 1500 LCD | Schott | Fiber optic light source | |
Xylocain Pumpspray | AstraZeneca GmbH | Lidocain, local anesthetic | |
Absorption Triangles – Unmounted | Fine Science Tools | 18105-03 | Absorption triangles for the surgery |
Parkell C&B Metabond clear powder L | Hofmeester dental | 013622 | Quick adhesive cement |
Parkell C&B Metabond Quick Base B | Hofmeester dental | 013621 | Quick adhesive cement |
Parkell C&B Metabond Universal Catalyst C | Hofmeester dental | 013620 | Quick adhesive cement |
Adjustable Precision Applicator Brushes | Parkell | S379 | Precision applicators for the surgery |
Blunt needles 0.9×23 mm | Dentina | 0441324 | Blunt needles |
Blunt needles 0.5×42 mm | Dentina | 0452155 | Blunt needles |
Blunt needles 0.3×23 mm | Dentina | 0553532 | Blunt needles |
Kallocryl A/C | Speiko | 1615 | Acrylic liquid component |
Kallocryl | Speiko | 1609 | Acrylic powder |
Hydrofilm transparent roll | Hartmann | Adhesive film | |
Head plates | Custom made | 30 mm x 10 mm size; 8 mm diameter hole, titanium | |
Head plate clamp | Custom made | Head plate holder | |
Pedestal post holders | Thorlabs | PH20E/M | Head plate holder |
Stainless steel post | Thorlabs | TR30/M | Head plate holder |
Stainless steel post | Thorlabs | TR75/M | Head plate holder |
Stainless steel post | Thorlabs | TR150/M | Head plate holder |
Post connector clamps | Custom made | Head plate holder | |
Aluminum Breadboard, 300 mm x 450 mm x 12.7 mm, M6 Taps | Thorlabs | MB3045/M | Microscope stage |
7" x 4" Lab Jack | Thorlabs | L490/M | Microscope stage |
Low profile face mask small mice | Emka Technologies | VetFlo-0801 | Anesthesia facemask holder |
RS4000 Tuned Damped Top Performance Optical Table | Newport | Floating table | |
S-2000A Top Performance Pneumatic Vibration Isolators with Automatic Re-Leveling | Newport | Floating table | |
Power Meter Model 1918-R | Newport | Power meter | |
X-Cite 120Q | Excelitas Technologies | Fluorescence lamp | |
Two-photon microscope | Bruker | Ultima IV | Two-photon microscopes |
Two-photon microscope | Thorlabs | Bergamo | Two-photon microscopes |
Plan N 4x/0.10 ∞/-/FN22 | Olympus | Objectives | |
Plan N 10x/0.25 ∞/-/FN22 | Olympus | Objectives | |
LMPlan FLN 20x/0.40 ∞/-/FN26.5 | Olympus | Objectives | |
XLPlan N 25x/1.00 SVMP ∞/0-0.23/FN18 | Olympus | Objectives | |
Ultafast tunable laser for 2P excitation | Spectraphysics | Mai Tai Deep See | Excitaiton lasers |
Ultafast tunable laser for 2P excitation | Spectraphysics | InSight DS+ Dual beam | Excitaiton lasers |
Ultafast tunable laser for 3P excitation | Coherent | Monaco | Excitaiton lasers |