We would like to thank all the members of the Molecular BioEngineering Group at Westlake University for all the help and useful discussion. We also thank Jinze Li and Jie-Min Jia from Westlake University for the help with filming the surgical procedure.
This work was supported by start-up funding from the Foundation of Westlake University, 2020 BBRF Young Investigator Grant, and National Natural Science Foundation of China grant 32050410298 all to K.D.P.

<ol>
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The authors have nothing to disclose.

Here we describe a method for long-term imaging of the hippocampus CA1 region in behaving mice. The method is based on the chronic implantation of a custom-made imaging window, which also enables the targeted administration of viruses or drugs directly to the neurons of interest. The present protocol consists of four major parts: i) assembling of imaging implant; ii) installation of imaging implant; iii) virus injection via imaging implant; iv) functional imaging in behaving mice. Below we describe and discuss critical steps in the protocol, troubleshooting, modifications, and limitations of the method. We also discuss the significance of the method and its potential alternative applications.
There are several critical steps in the described protocol that are rather important for successful surgery: (i) preparation of a high-quality imaging implant; (ii) sterile surgical conditions; (iii) aspiration of the cortex; (iv) precise placing of the imaging implant; (v) viral injection. As Step 1.6 indicates, excess soldering tin would require a larger craniotomy and thus increases the risk of inflammation. It is also very important to use an appropriate amount of the adhesive optical glue when fixing the cover glass to the imaging cannula, as indicated in Step 1.11, as an insufficient amount may result in leakage of cerebrospinal fluid into the imaging cannula and making it opaque. On the other hand, excess optical adhesive can result in the decreased transparency of the glass window. Possible contamination of the imaging implant can cause an active proliferation of connective tissue under the optical window and/or severe inflammation, which will lead to early termination of the experiment. Therefore, imaging implant assembling and preparation before the surgery is almost as important as the surgical procedure itself.
During the surgery, the part of the cortex under craniotomy is ablated by gentle aspiration, which results in inevitable bleeding. Blood at the surgical site significantly reduces the visibility of the brain tissue that must be removed. This complicates the precise assessment of the required depth of tissue ablation. Careful flushing of the surgical site with PBS every time before applying suction to remove the next portion of tissue provides better control of depth. The brain tissue should always be removed in small portions step-by-step confirming the depth of ablated tissue before proceeding with more suction. Finer control of suction can also be achieved with a thinner blunt needle. We suggest using a 26 G needle, however, smaller than 26 G diameter is more prone to clogging. Furthermore, it usually takes lots of practice to determine the precise depth of aspiration required for each animal, as the color of the cortex, corpus callosum, and hippocampus may vary from mouse to mouse (Figure 3).
Insertion and securing of the imaging implant should be done very precisely to ensure the closest possible position of the imaging window to the dorsal surface of the hippocampus. The size of the prepared craniotomy should closely match the implant and allow its insertion without significant resistance. At the same time, there should be no visible gap between the skull and the implant to ensure proper sealing and avoid brain tissue exposure. Gentle and stable pressure should be applied to the top of the implant during its sealing to the skull. It is almost unavoidable to have blood under the imaging window during the implant installation. If the surgical procedure is done properly, the window should clear out in 3-7 days, and brain vasculature becomes clearly visible. It is also important to ensure that virus is properly injected under the window. In the case of failed expression, virus can be reinjected multiple times.
The major complication we encountered in some cases is reduced visibility of the imaging window. There are several possible reasons for poor imaging quality: i) ongoing inflammation; ii) outgrowth of connective tissue on the glass; iii) big gap between the window and hippocampus. Inflammation is usually caused by contamination during surgery or by not properly sterilized imaging implant. We suggest autoclaving the surgical instruments before and after each surgery, disinfect the surgery area right before the procedure, and wear clean personal protective equipment during surgery. Imaging implants should be cleaned after assembling, sterilized, and stored in sterile conditions. The outgrowth of connective tissue on the glass of imaging implant can be due to mechanical contamination on the surface of the glass or excessive trauma of the brain tissue during cortex ablation. After assembling the implant, it is important to confirm the glass surface is clean and smooth. Also, all pieces of damaged brain tissue must be carefully removed before inserting imaging implant into the craniotomy. In certain cases, the gap between the glass window and hippocampus results in the accumulation of cerebrospinal fluid, reducing the quality of imaging. Therefore, during implant installation, it is crucial to insert it all the way in ensuring good contact between hippocampus and glass window. Sometimes, it is difficult to identify the exact reason for opaque imaging window. We suggest performing post-mortem analysis to reveal conditions under the optical window and correspondingly adjust subsequent surgeries.
The method has several fundamental and technical limitations that should be taken into account before and during in vivo imaging. One of the major limitations is cortex ablation. Part of the visual and sensory cortex is removed during the surgery. While it is hard to precisely evaluate the impact of cortex ablation, as removed brain tissue does not directly project onto hippocampus, several studies demonstrated no noticeable impairment of hippocampal-dependent learning or other relevant hippocampus functions15,16. The optical limitations should also be considered, especially when high NA objective lenses are used. For example, in this study, we used a 1.75 mm long cannula with a 1.9 mm inner diameter. The geometry of this cannula will not preserve the full NA of air objective with NA more than ~0.5 or water objective with NA more than ~0.6 as it will clip some of light. Another limitation, common for all brain imaging implants, is that part of the brain is getting exposed, thus promoting heat loss17,18. However, physiological brain temperature can be easily restored during imaging by perfusion of a warm buffer.
The described method can be easily modified or adjusted for other applications. For example, the preparation can be adapted for imaging of striatum7. As the striatum lays slightly deeper than hippocampus, the longer imaging cannula should be used for assembling imaging implant. We suggest using 2.0 mm imaging cannula. The coordinates of the craniotomy should be adjusted correspondingly (AP: +0.8 mm, ML: &#8722;1.8 mm). In addition, virus injection via infusion cannula allows achieving the expression of a transgene in a thin layer of neurons when using AAV serotype with restricted spread19,20. It is particularly beneficial for one-photon imaging due to reduced out-of-focus fluorescence from deeper layers and, as a result, improved single-cell resolution imaging. Furthermore, injection cannula can also be used during functional imaging for the administration of drugs or other chemicals directly onto the neurons in FOV (Figure 5B). Overall infusion cannula adds useful functionalities to the imaging implant, improving imaging quality due to the targeted viral expression and enabling pharmacological stimulation of neurons in FOV. The used head plate provides extraordinary stability of imaging implant minimizing motion artifacts even in actively moving animals on a treadmill. The head plate is small and light, causing minimal discomfort to animals, and remains stable for several months after installation. The imaging implant is also compatible with multiphoton imaging15,16,21 and can be combined with micro-endoscopes22,23. A similar imaging implant was also used for multiphoton imaging of the deeper hippocampus structures, including stratum radiatum, stratum lacunose, and dentate gyrus16,24,25,26,27. However, targeting the deeper hippocampus structures with AAVs via infusion cannula may require further optimization of the AAV serotype and volume19.
We believe that the described protocol will facilitate studies that aim to investigate neuronal activity with high spatiotemporal resolution in the hippocampus of behaving mice using simple and affordable one-photon imaging setups.

The hippocampus is a key brain region responsible for learning and memory1 as well as for spatial navigation2. Hippocampal atrophy is associated with neurological and psychiatric disorders involving memory loss and cognitive decline3,4,5. In mice, the hippocampus is a very well-established model to study spatial, contextual, and associative learning and memory formation on the cellular and network levels4,5. The mechanistic studies of learning and memory require longitudinal interrogation of neuronal structure and function in behaving mice. Fluorescence imaging in combination with genetically encoded probes6 provides unprecedented capabilities to record membrane voltage dynamics7,8, calcium transients9, and structural changes10 over large subsets of neurons intravitally. However, optical access to the hippocampus in mice is obstructed by the cortex, which can reach over 1 mm in thickness. Here, we described a procedure for assembling a custom-made imaging device and its chronic implantation into the mouse head for long-term optical access to the CA1 subregion of the dorsal hippocampus in behaving mice. Infusion cannula integrated into the imaging implant allows administration of viruses or drugs directly onto the neurons in the field of view. The described preparation in combination with wide-field microscopy enables recurrent imaging of the large subsets of neurons in behaving mice over long periods of time. We utilized this preparation to express calcium and voltage genetically encoded indicators in hippocampal CA1 region via targeted injection of recombinant adeno-associated virus (rAAV) for neuronal activity recordings at single-cell resolution. We also performed longitudinal calcium imaging of the corresponding neuronal subsets at high spatiotemporal resolution in behaving animals. In addition, this preparation is compatible with multiphoton microscopy and microendoscopy, thus further expanding the toolbox of imaging techniques to study neuronal networks at cellular and subcellular levels in behaving mice. We described critical steps and troubleshooting of the protocol. We also discussed the possible pitfalls and limitations of the method.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cover glass</td>
			<td>Deckgl&#228;ser</td>
			<td>72296-03</td>
			<td></td>
		</tr>
		<tr>
			<td>denture&#160;base resin</td>
			<td>ShangHai New Centery Dentel Material</td>
			<td>N/A</td>
			<td>Type I, self-solidifying</td>
		</tr>
		<tr>
			<td>Dummy Cannula</td>
			<td>RWD Life Science Co.,LTD</td>
			<td>62102</td>
			<td>OD 0.30mm</td>
		</tr>
		<tr>
			<td>Guide Cannula</td>
			<td>RWD Life Science Co.,LTD</td>
			<td>62003</td>
			<td>26G</td>
		</tr>
		<tr>
			<td>Head Fork</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom made</td>
		</tr>
		<tr>
			<td>Head Plate</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom made</td>
		</tr>
		<tr>
			<td>Imaging Cannula</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom Made; OD 3mm, ID 2.7mm, Height 1.8mm, #108 stainless steel</td>
		</tr>
		<tr>
			<td>Internal Cannula</td>
			<td>RWD Life Science Co.,LTD</td>
			<td>62203</td>
			<td>&#160;0.30*0.14 (OD*ID, mm)</td>
		</tr>
		<tr>
			<td>Kwik Sil</td>
			<td>World Precision Instruments</td>
			<td>KWIK-SIL</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperBond C&#38;B</td>
			<td>SUN MEDICAL</td>
			<td>N/A</td>
			<td>SuperBond C&#38;B kit</td>
		</tr>
		<tr>
			<td>Treadmill Kit</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom made</td>
		</tr>
		<tr>
			<td>UV-cured Adhesive</td>
			<td>NORLAND PRODUCTS</td>
			<td>NOA 60</td>
			<td></td>
		</tr>
	</tbody>
</table>

craniotomy procedure for visualizing neuronal activities in hippocampus of behaving mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

In vivo imaging of neuronal activity using a genetically encoded calcium indicator. On average, in vivo imaging starts 3-4 weeks after implantation if a sufficient level of transgene expression is achieved. By this time, cerebral edema and hemorrhage are usually completely resolved, and brain vasculature can be easily observed through the optical window. Here we utilized the described preparation to perform the repeated recordings of neuronal activity in the dorsal hippocampal CA1 region in behaving mice under fluorescence wide-field microscope. To record neuronal activity, we used a bright genetically encoded calcium indicator, named NCaMP713, which exhibits similar calcium sensitivity and temporal resolution to that of GCaMP6s14. To express the NCaMP7 indicator in the hippocampus, we injected the rAAV/DJ-CAG-NCaMP7 virus using an infusion cannula and initiated longitude imaging at 14 days post-injection. To record neuronal activity, we used a 10x NA 0.3 air objective lens and Hamamatsu OrcaFusion sCMOS camera that allowed imaging at ~1.5x1.5 mm FOV at up to 100 Hz frequency. Green fluorescence was excited by a commercially available 470 nm LED using a standard GFP filter set. The average depth of imaging achieved in green channel is about 50-120 &#181;m, which allows recording of neuronal activity mainly in stratum oriens and stratum pyramidale. Imaging depth in near-infrared channels can be up to 200 &#181;m reaching the deeper layers of hippocampus8. The average recording time per FOV was 6-12 min, although much longer imaging sessions are possible as NCaMP7 is characterized by extremely high photostability and no detectable phototoxicity was observed (Figure 6).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62266/62266fig06.jpg" /> 
Figure 6: Recording of the neuronal activity in the hippocampal neurons using a green fluorescence genetically encoded calcium indicator. (A) A selected FOV imaged under a wide-field fluorescence microscope in green channel. (B) The 15 ROIs corresponding to the single neurons shown in A and selected using the standard deviation projection of the whole recording. (C) Representative fluorescence single-trial traces of the 15 selected neurons in B. (D) A representative zoom-in view of 2 calcium traces shown in corresponding color boxes shown in C. Scale bar, 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62266/62266fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To obtain fluorescence traces, the regions of interest (ROIs) corresponding to neuronal somas were segmented manually and analyzed by ImageJ software. Prior to the image analysis motion correction, common post-recording procedures in awake animals were not required as the acquired datasets did not exhibit motion artifacts due to the high stability of imaging implant. A representative single-trial optical recording of neuronal activities from the hippocampus in an awake behaving mouse is presented in Figure 6. 15 ROIs corresponding to neuronal somas were selected manually from the same FOV shown in Figure 6B, and the single-trial fluorescence traces within each ROI are shown in Figure 6C. Figure 6D shows two representative parts of fluorescence traces from two different ROIs. We performed 4 consecutive imaging sessions for the same FOV with 3-day intervals. It was possible to identify and image the same neurons in certain FOVs for at least two weeks (the longer imaging sessions were not performed for this study, however, the same preparation has been used for up to 6 month-long imaging study in mice previously7; Figure 7). In this study, we used AAV/DJ-CAG vector, which was driving a strong expression of the gene of interest even 21 days after virus delivery (Supplementary Figure 1). The continuous expression complicated long-term identification of the same neurons due to increased fluorescence background and appearance of new neurons expressing the calcium indicator. Therefore, selection of the AAV serotype and promoter to drive target gene expression should be one of the important considerations during experimental design in particular if longitudinal imaging of the same subset of neurons is required. The imaging quality allowed to resolve proximal dendrites as well as visualize blood vessels.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62266/62266fig07v2.jpg" /> 
Figure 7: Image sequence of four different FOVs from the hippocampal area tracked over 12 days. The animal was implanted with the window on Day 0 and was injected with the rAAV/DJ-CAG-NCaMP7 virus on Day 7. Arrowheads indicate the neuron tracked within the FOV. Scale bars: 80 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62266/62266fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In vivo imaging of neuronal activity using a genetically encoded voltage sensor. 
In this study, we also used a novel genetically encoded voltage sensor, named SomArchon7, which allows for voltage imaging with single-cell single-spike resolution in behaving animals7,8. To express SomArchon, we injected rAAV/DJ-CAG-SomArchon virus using an infusion cannula and performed voltage imaging in a head-fixed behaving mouse several days post-injection. To record neuronal activity, we used a 40x NA 0.8 objective lens and Hamamatsu OrcaFusion sCMOS camera that allowed us to image 150x40 &#181;m FOV at up to 830 Hz acquisition rate. The GFP protein, which a part of SomArchon construct to facilitate visualization of expression in the visible range of the spectrum, can be easily imaged in the green channel (LED excitation at 470/20 nm, emission 525/50 nm) to locate cells of interest for voltage imaging. Optical voltage recordings were performed in the near-infrared channel (laser excitation 637 nm at 3.4 W/mm2, emission 665 nm long pass) with 4 x 4 binning at 830 Hz acquisition rate. We recorded the spontaneous activity of a hippocampal neuron in an awake mouse with average SNR of 7 per action potential (Figure 8). 
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/62266/62266fig08.jpg" /> 
Figure 8: Recording of the neuronal activity in the hippocampal neurons using a near-infrared fluorescence genetically encoded voltage indicator SomArchon. (A) A selected FOV imaged under wide-field fluorescence microscope in the near-infrared channel. (B) Single-trial fluorescence trace of the neuron in A. (C) A representative zoom-in view of the voltage trace in the corresponding box shown in B. Scale bar: 25 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62266/62266fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Histology 
After functional imaging study is done, post-mortem analysis is used to confirm the correct placement of the implant, the area of virus expression, and localization of a protein of interest in imaged neurons. For histological verification of virus expression and placement of the implant, coronal sections of the PFA fixed brain were examined under a fluorescence wide-field microscope (Figure 9A, C). A confocal microscope was used to acquire high-resolution images of individual neurons expressing the calcium indicator, as well as the voltage indicator (Figure 9B, D). DAPI staining was used to visualize the overall morphology of the brain slice. In addition, the brain slices can be assessed using immunohistochemistry to verify astrogliosis or gliosis caused by window implantation and viral expression. Our previous studies demonstrated that the procedure did not induce noticeable gliosis7.
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/62266/62266fig09.jpg" /> 
Figure 9:&#160;Histological verification of the optical window position and virus expression. (A) A representative fluorescent image of the coronal section brain slice showing the placement of the optical window from an NCaMP-expressing mouse. Scale bar: 1 mm. (B) A representative confocal image of neurons expressing the NCaMP7 indicators. Scale bar: 25 &#181;m. (C) A representative fluorescence image of the coronal section brain slice showing the placement of the optical window from a SomArchon-expressing mouse. Scale bar: 1 mm. (D) A representative confocal image of neurons expressing the SomArchon indicators. Scale bar: 25 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62266/62266fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Quantitive analysis of relative fluorescence intensity along with expression time. <a href="https://www.jove.com/files/ftp_upload/62266/Supplementary Figure 1.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice at JoVE.com

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

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7.0K Views.  Westlake University. This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice. 

Video: Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2 connecting the AL with the MB.&#160;The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi&#160;channel wire electrodes is demonstrated. Pairwise differential amplification of the micro&#160;wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long&#160;term recordings over many hours up to days, which is a clear advantage compared to conventional extra&#160;and intracellular in vivo recording techniques.

Simultaneous extracellular long term recordings from two different brain neuropiles or two different anatomical tracts were established in honeybees. These recordings allow the investigation of temporal aspects of neuronal processing across different brain areas at the single neuron as well as at the ensemble level in a behaving animal.

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent ...

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, typically 3 mm x 3 mm. This protocol introduces a method for creating a substantially larger 7 mm x 6 mm cranial window exposing most of a cerebral hemisphere over the mouse temporal and parietal cortices (e.g., bregma 2.5 - 4.5 mm, lateral 0 - 6 mm). To perform this surgery, the head must be tilted approximately 30&#176; and much of the temporal muscle must be retracted. Due to the large amount of bone removal, this procedure is intended only for acute experiments with the animal anesthetized throughout the surgery and experiment.
The main advantage of this innovative large lateral cranial window is to provide simultaneous access to both medial and lateral areas of the cortex. This large unilateral cranial window can be used to study the neural dynamics between cells, as well as between different cortical areas by combining multi-electrode electrophysiological recordings, imaging of neuronal activity (e.g., intrinsic or extrinsic imaging), and optogenetic stimulation. Additionally, this large craniotomy also exposes a large area of cortical blood vessels, allowing for direct manipulation of the lateral cortical vasculature.

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Long-term Sensory Conflict in Freely Behaving Mice

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning in mice. By permanently wearing a device fixed on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages. Therefore, this protocol readily enables the study of the visual system and multisensory interactions over an extended timeframe that would not be accessible otherwise. In addition to lowering the experimental costs of long-term sensory learning in naturally behaving mice, this approach accommodates the combination of in vivo and in vitro experiments. In the reported example, video-oculography is performed to quantify the vestibulo-ocular reflex (VOR) and optokinetic reflex (OKR) before and after learning. Mice exposed to this long-term sensory conflict between visual and vestibular inputs presented a strong VOR gain decrease but exhibited few OKR changes. Detailed steps of device assembly, animal care, and reflex measurements are hereby reported.

The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning. By permanently wearing a fixed device on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages.

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for ...

Visualizing Protein Kinase A Activity In Head-fixed Behaving Mice Using In Vivo Two-photon Fluorescence Lifetime Imaging Microscopy

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. Despite their importance, technologies for tracking neuromodulatory events with cellular resolution are just beginning to emerge. Neuromodulators, such as dopamine, norepinephrine, acetylcholine, and serotonin, trigger intracellular signaling events via their respective G protein-coupled receptors to modulate neuronal excitability, synaptic communications, and other neuronal functions, thereby regulating information processing in the neuronal network. The above mentioned neuromodulators converge onto the cAMP/protein kinase A (PKA) pathway. Therefore, in vivo PKA imaging with single-cell resolution was developed as a readout for neuromodulatory events in a manner analogous to calcium imaging for neuronal electrical activities. Herein, a method is presented to visualize PKA activity at the level of individual neurons in the cortex of head-fixed behaving mice. To do so, an improved A-kinase activity reporter (AKAR), called tAKAR&#945;, is used, which is based on F&#246;rster resonance energy transfer (FRET). This genetically-encoded PKA sensor is introduced into the motor cortex via in utero electroporation (IUE) of DNA plasmids, or stereotaxic injection of adeno-associated virus (AAV). FRET changes are imaged using two-photon fluorescence lifetime imaging microscopy (2pFLIM), which offers advantages over ratiometric FRET measurements for quantifying FRET signal in light-scattering brain tissue. To study PKA activities during enforced locomotion, tAKAR&#945; is imaged through a chronic cranial window above the cortex of awake, head-fixed mice, which run or rest on a speed-controlled motorized treadmill. This imaging approach will be applicable to many other brain regions to study corresponding behavior-induced PKA activities and to other FLIM-based sensors for in vivo imaging.

A procedure is presented to visualize protein kinase A activities in head-fixed, behaving mice. An improved A-kinase activity reporter, tAKAR&#945;, is expressed in cortical neurons and made accessible for imaging through a cranial window. Two-photon fluorescence lifetime imaging microscopy is used to visualize PKA activities in vivo during enforced locomotion.

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.