We gratefully acknowledge support by the Harold and Pauline Price Chair in Ophthalmology and the Jules Stein Eye Institute to MBG, the NEI Core grant (EY00331-43) to SN. Research was supported in part by a generous gift from the Sakaria family to SN and MGB, and from an unrestricted grant from the Research to Prevent Blindness to the Department of Ophthalmology. We thank Charlotte Yiyi Wang at Berkeley School of Optometry for obtaining initial OCT images of subretinal injections.

Animals: Wild type C57Bl/6J mice bred at the University of California at Los Angeles (UCLA). All animals were between 11 - 17 weeks old, and included male and female mice. All mice were group-housed, maintained in a 12:12 light/dark cycle with food and water ad libitum. All experiments were performed in accordance with the institutional guidelines of UCLA and the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research.
NOTE: All drugs and injectable agents are United States Pharmacopeia (USP) grade.
1. Surgical Preparation
<ol>
	<li>Anesthetize the mouse with an intraperitoneal injection of 100 mg/kg ketamine and 8 mg/kg xylazine in a saline mix. Administer anesthesia to a depth such that the mouse has no toe pinch or corneal touch reflexes.</li>
	<li>Maintain body temperature at 37.0 &#176;C with a circulating water pad.</li>
	<li>Dilate pupils with 2.5% phenylephrine eye drops and trim whiskers to facilitate visualization. Whiskers provide significant sensory input to the mouse, therefore, whisker trimming should remove only the portion that blocks clear access to the eye, and not to the base of the whisker. In our experience, mice show normal recovery after this procedure.&#160;Apply methylcellulose eye drops to prevent dryness and minimize anesthetic induced transient cataracts13.</li>
	<li>Sterilize instruments prior to surgery (i.e., betadine and ethanol or hot beads).</li>
	<li>Prepare diluted fluorescein (0.01% using 0.9% saline) in a sterile environment (i.e., biosafety cabinet) if visualization will be performed (see section 3 below).</li>
</ol>
2. Injection Site Preparation
<ol>
	<li>Prepare a syringe (e.g., 5 &#181;l syringe) with the appropriate injection volume (e.g., 0.3 to 1.0 &#181;l).</li>
	<li>Position the mouse so the eye is facing up and clearly visible in the dissecting microscope.</li>
	<li>Gently pinch the temporal conjunctiva with fine tipped forceps. Make a circumferential incision of approximately 90 degrees using curved Vannas scissors.</li>
	<li>Repeat step 2.3 with the underlying Tenon&#39;s capsule.</li>
	<li>Resect the surrounding connective tissue with fine tipped forceps while rotating the globe nasally. Work towards the injection site approximately 0.5 mm temporal to the optic nerve. Use great care to avoid disrupting the retro-orbital sinus.</li>
</ol>
3. Sclerotomy and Subretinal Injection
NOTE: It is recommended that the injection of 0.01% fluorescein in 0.9% Saline be used to assist with visualization while learning this procedure. The topographic distribution of fluorescein can be effectively documented with fundus imaging (see section 4 below).
<ol>
	<li>Make a small scleral incision at the injection site by gently scratching the eyecup with a 22.5-degree ophthalmic blade. This incision should only be large enough to allow the tip of the needle to pass through the sclera.</li>
	<li>Insert the beveled 33 G needle (angled 5 - 10&#176; into the sclerotomy with the bevel facing and angled parallel to the retina. Inject desired volume (e.g., 0.3 to 1.0 &#181;l of 0.01% Fluorescein for learning purposes). 
	NOTE: Maintain sterility of the syringe by thoroughly cleaning with successive washes of a suitable solvent and DI water before each injection.</li>
	<li>Depress the plunger slowly (over ~ 3 sec) without moving the needle and with even pressure. 
	NOTE: When the needle is in the subretinal space, a slight resistance will be felt while depressing the plunger. There will be no to minimal resistance if the needle punctures the retina, and high resistance if the needle does not penetrate the sclera or RPE.</li>
	<li>Wait several seconds before withdrawing the needle to minimize backflow.</li>
	<li>Rinse the eye with sterile buffered saline and ensure the eye has rotated back to its normal position.</li>
</ol>
4. Assessment of Retinal Detachment by OCT and Fundus Imaging
<ol>
	<li>Perform OCT imaging immediately after injection to evaluate the quality of the injection and at appropriate time points post-injection as needed to evaluate retinal structure. 
	NOTE: Examples of the use of OCT in similar studies has been previously described7,14.
	<ol>
		<li>Adjust and align the OCT image to target the site of injection. The injection site should be midline and 0.5 mm temporal to the optic nerve head. Repeat as needed if the detachment is out of frame or not optimally centered.</li>
	</ol>
	</li>
	<li>Visualize retinal detachment and dye injection area with en-face fundus imaging7,14. 
	NOTE: If an OCT imaging system is not available, injection of a small amount of fluorescein with a vector for practice will allow visualization with any fundus camera that performs fluorescein angiography using the same excitation wavelengths and blocking filters. Localized areas of hyper-fluorescence will appear underneath the vasculature and the vasculature will have sharp and distinct boundaries if the subretinal space is targeted correctly. The edge of the bleb from the injection will be demarcated by the transition from hyper- to hypo- fluorescence. Several instruments provide this capability for the mouse; the instrumentation used here is described elsewhere14.</li>
</ol>
5. Post-operative Care
<ol>
	<li>Apply a thick coat of triple antibiotic ophthalmic cream to the corneal surface of the injected eye.</li>
	<li>Place mice in clean solitary cages for recovery. Do not combine mice that have undergone surgery until they are fully recovered.</li>
	<li>Monitor respiration and temperature during anesthesia recovery. Monitor animals until they can maintain sternal recumbency.</li>
	<li>Perform additional appropriate post-operative monitoring and treatment, including a sub-cutaneous injection of carprofen (5 mg/kg) for post-surgical pain management.</li>
</ol>
6. Assessment of Retinal Function by Electroretinography (ERG)
<ol>
	<li>Perform ERG analysis pre-injection and at appropriate times post-injection as needed to evaluate retinal function. If the injection was made into the subretinal space, the retinal detachment should resolve within 72 hr.
	<ol>
		<li>Use standard ERG techniques to evaluate retinal function before and after injection as previously described14,15.</li>
	</ol>
	</li>
</ol>
7. 3D Reconstruction and Bleb Volume Quantification
NOTE: OCT scans with high contrast encompassing the entire detachment within the frame of view are optimal for use. ImageJ/Fiji17,18 and Imaris were used, but other software can be used.
<ol>
	<li>Export the b-scan of interest, import to ImageJ/Fiji and crop (Image &#62; Crop) the portion of the OCT scan to be modeled using the rectangular selection tool.
	<ol>
		<li>Adjust contrast (Image &#62; Adjust &#62; Brightness/Contrast) and delineate any missing boundaries by connecting two sections with a line.</li>
		<li>Draw a straight line with the line tool (holding shift) that spans the RPE to the photoreceptor layer. Measure (Analyze &#62; Measure) the length of the line to obtain size of maximum detachment for step 7.8.</li>
	</ol>
	</li>
	<li>Import cropped frames to the 3D-reconstruction software (See Table of Materials) using the &#34;RGB to Gray&#34; plugin and MATLAB Compiler Runtime.</li>
	<li>Set the voxel size (under Image Properties) using the calibration parameters from the OCT scan (x,y,z).</li>
	<li>Execute the &#34;RGB to Gray&#34; plugin (under Image Properties), with equal weighting to each channel, to create a fourth channel. Delete original Red-Green-Blue channels.</li>
	<li>Invert the gray channel using contrast change. Store Image.</li>
	<li>Click the &#34;add a new surface&#34; button in the 3D-View tab, and begin the guided 4 step process of creating the surface.
	<ol>
		<li>Set the surface level detail (step 1 of 4). 
		NOTE: In our experience 8.0 to 12.0 was the most effective range.</li>
		<li>Set the maximum sphere size (under Background Selection) to slightly less than the maximum detachment size measured in 7.1.2. Create the surface and undo gray channel inversion (step 2 of 4).</li>
		<li>Set the threshold to the maximum value so the surface of the negative spaces outside of the retina and the detachment do not come in contact (step 3 of 4).</li>
		<li>Set the filter type to the number of voxels and isolate the negative space in the detachment site by size. Finish the surface (step 4 of 4). 
		NOTE: The volume of the detachment surface is located under volume in the statistics tab.</li>
	</ol>
	</li>
</ol>

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None of the authors have any commercial disclosures.

Subretinal injections are the method of choice for the delivery of viral vectors and stem cell-derived therapy for manipulating photoreceptors and the RPE in both basic research and clinical treatment. In patients, subretinal injections are typically done with an anterior sclerotomy at the pars plana, a posterior core vitrectomy and penetration of the retina by the needle with direct visualization. As with most vitrectomy procedures, it is common for cataract formation to occur prematurely unless the eye is already pseud...

Subretinal injections are the primary means of delivering cellular and viral agents to the retinas of mice to study their effects on photoreceptors and the underlying RPE1,2. Most subretinal injection protocols in mice use a transcorneal or a transcleral injection site anterior to the equator (Figure 1). This approach can result in inherent collateral damage that includes nicking and resultant clouding of the lens, disruption of the integrity of the vitreous, penetration of the neurosensory retina and iris, retinal hemorrhage, substantial retinal detachments and lasting subretinal edema 3-9. Experimental manipulations must overcome these effects in order to evaluate the effects of therapeutic interventions3,7,10,11. This study provides a detailed description and validation of a posterior transcleral injection method that avoids these complications, minimizes trauma and has a high success rate of targeting the subretinal space.
Injections targeting the subretinal space in mice are often very difficult to perform and most investigators encounter a high frequency of failed attempts in which the vector is delivered to an incorrect location or there is significant retinal damage, for example in a complete retinal detachment6. The number of eyes excluded from analysis because of injection complications is typically not reported in mouse studies, but in our own experience and in discussion with other investigators, the number of failed injections can be as high as 50% and vary dependent on the experience and capabilities of the investigator who is performing the injections. The success of the injection is typically assessed by direct fundus imaging and/or optical coherence tomography (OCT)7,9. An easily mastered method with high success rates for subretinal injections in mice can hasten experimentation and reduce the cost of preclinical studies of treatments for retinal diseases that are major causes of blindness in the United States.
The posterior, transcleral subretinal injection technique described here is an adaptation from clinical and preclinical protocols9,12. The noninvasive diagnostic assessments performed in injected mice demonstrate mild and highly localized damage and lack additional collateral lens, retinal and RPE injury. Furthermore, with relatively little practice, an experimenter can achieve these results with a high success rate (80 - 90% or better), thereby reducing the costs associated with such studies. This procedure can be used to deliver cellular, viral, or pharmacological therapeutic interventions to photoreceptors and/or RPE in preclinical studies and to easily evaluate experimental interventions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Hamilton Model 62 RN SYR</td>
			<td>Hamilton</td>
			<td>87942</td>
			<td>Syringe x 1</td>
		</tr>
		<tr>
			<td>Hamilton Needle 33G, 1.0&#34;, 20 DEG, point 3 (304 stainless steel)</td>
			<td>Hamilton</td>
			<td>7803-05</td>
			<td>Needles x 6</td>
		</tr>
		<tr>
			<td>Vannas Curved Scissors</td>
			<td>Ted Pella, INC.</td>
			<td>1347</td>
			<td>5mm Blade</td>
		</tr>
		<tr>
			<td>22.5 Degree Microsurgery Knife</td>
			<td>Wilson Ophthalmic Corp.</td>
			<td>91204</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketaject&#160;</td>
			<td>Phoenix</td>
			<td>NDC 57319-609-02</td>
			<td>Ketamine</td>
		</tr>
		<tr>
			<td>Anased</td>
			<td>Lloyd Laboratories</td>
			<td>NDC 61311-482-10</td>
			<td>Xylazine</td>
		</tr>
		<tr>
			<td>Fluorescein 10% AK-Fluor</td>
			<td>Akorn</td>
			<td>NDC 17478-253-10</td>
			<td>100mg/ml</td>
		</tr>
		<tr>
			<td>0.9% Saline USP</td>
			<td>Hospira</td>
			<td>NDC 0409-4888-50</td>
			<td>0.9% NaCl</td>
		</tr>
		<tr>
			<td>Antibiotic Ointment</td>
			<td>Akorn</td>
			<td>NDC 17478-235-35</td>
			<td>Ophthalmic</td>
		</tr>
		<tr>
			<td>Water Circulating Pump</td>
			<td>Gaymar</td>
			<td>TP-500 T/Pump&#160;</td>
			<td>P/N 07999-000</td>
		</tr>
		<tr>
			<td>sd-OCT</td>
			<td>Bioptigen</td>
			<td>R-Series</td>
			<td>Commercial</td>
		</tr>
		<tr>
			<td>Fundus Camera</td>
			<td>Phoenix Research Laboratories</td>
			<td>MICRON III</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers Type 3</td>
			<td>Ted Pella, INC.</td>
			<td>5385-3SU</td>
			<td></td>
		</tr>
		<tr>
			<td>2.5% Phenylephrine</td>
			<td>Paragon BioTeck</td>
			<td>NDC 42702-102-15</td>
			<td>Ophthalmic</td>
		</tr>
		<tr>
			<td>IMARIS8</td>
			<td>Bitplane</td>
			<td></td>
			<td>Version 8.1.2</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td>V1.8.0_77</td>
		</tr>
		<tr>
			<td>Hypromellose&#160;2.5%</td>
			<td>Goniovisc</td>
			<td>AX0401</td>
			<td>Methylcellulose</td>
		</tr>
		<tr>
			<td>Eye Drops (Rinse)</td>
			<td>Bausch &#38; Lomb</td>
			<td></td>
			<td>Saline Solution</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Zeiss</td>
			<td>Stemi 2000</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Light source</td>
			<td>Fostec</td>
			<td>P/N 20520</td>
			<td>Light source</td>
		</tr>
	</tbody>
</table>

an alternative and validated injection method for accessing the subretinal space via a transcleral posterior approach

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to the interphotoreceptor/subretinal compartment that has direct exposure to photoreceptors and the retinal pigment epithelium (RPE). Subretinal injections of plasminogen as well as recent preclinical and clinical trials have demonstrated safety and/or efficacy of delivering viral vectors and stem cells to individuals with advanced retinal disease. Mouse models of retinal disease, particularly hereditary retinal dystrophies, are essential for testing these therapies. The most common injection procedure in rodents is to use small transcorneal or transcleral incisions with an anterior approach to the retina. With this approach, the injection needle penetrates the neurosensory retina disrupting the underlying RPE and on insertion can easily nick the lens, causing lens opacification and impairment of noninvasive imaging. Accessing the subretinal space via a transcleral, posterior approach avoids these problems: the needle crosses the sclera approximately 0.5 mm from the optic nerve, without retinal penetration and avoids disrupting the vitreous. Collateral damage is limited to that associated with the focal sclerotomy and the effects of a transient, serous retinal detachment. The simplicity of the method minimizes ocular injury, ensures rapid retinal reattachment and recovery, and has a low failure rate. The minimal damage to the retina and RPE allows for clear assessment of the efficacy and direct effects of the therapeutic agents themselves. This manuscript describes a novel subretinal injection technique that can be used to target viral vectors, pharmacological agents, stem cells or induced pluripotent stem (iPS) cells to the subretinal space in mice with high efficacy, minimal damage, and fast recovery.

Posterior approach transcleral subretinal injections were performed on 31 healthy eyes from 16 wild type mice with injections of 0.3 &#181;l (n = 18), 0.5 &#181;l (n = 8) and 1.0 &#181;l (n = 5) of 0.01% fluorescein. One eye was excluded from injection due to a pre-existing corneal opacity that prevented structural and functional analysis. Every injected eye is included in this report. No unintended retinal detachments, punctures of the neurosensory retina, or leakage into the vitreous we...

Watch this Scientific Journal Video about An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach at JoVE.com

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to ...

an-alternative-validated-injection-method-for-accessing-subretinal

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Neuroscience

13.1K Views.  University of California, Los Angeles. Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

Video: An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Novel Apparatus and Method for Drug Reinforcement

Animal models of reinforcement have proven to be useful for understanding the neurobiological mechanisms underlying drug addiction. Operant drug self-administration and conditioned place preference (CPP) procedures are expansively used in animal research to model various components of drug reinforcement, consumption, and addiction in humans. For this study, we used a novel approach to studying drug reinforcement in rats by combining traditional CPP and self-administration methodologies. We assembled an apparatus using two Med Associate operant chambers, sensory stimuli, and a Plexiglas-constructed neutral zone. These modifications allowed our experiments to encompass motivational aspects of drug intake through self-administration and drug-free assessment of drug/cue conditioning strength with the CPP test. In our experiments, rats self-administered cocaine (0.75 mg/kg/inj, i.v.) during either four (e.g., the "short-term") or eight (e.g., the "long-term") alternating-day sessions in an operant environment containing distinctive sensory cues (e.g., olfactory and visual). On the alternate days, in the other (differently-cued) operant environment, saline was available for self-infusion (0.1 ml, i.v.). Twenty-four hours after the last self-administration/cue-pairing session, a CPP test was conducted. Consistent with typical CPP findings, there was a significant preference for the chamber associated with cocaine self-administration. In addition, in animals undergoing the long-term experiment, a significant positive correlation between CPP magnitude and the number of cocaine-reinforced lever responses. In conclusion, this apparatus and approach is time and cost effective, can be used to examine a wide array of topics pertaining to drug abuse, and provides more flexibility in experimental design than CPP or self-administration methods alone.

Operant drug self-administration and conditioned place preference (CPP) procedures are expansively used in research to model various components of drug reinforcement, consumption, and addiction in humans. In this report, we combined traditional CPP and self-administration methods as a novel approach to studying drug reinforcement and addiction in rats.

Animal models of reinforcement have proven to be useful for understanding the neurobiological mechanisms underlying drug addiction.  Operant drug ...

A Method of Nodose Ganglia Injection in Sprague-Dawley Rat

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.

Afferent vagal signaling transmits important information to central nervous system from receptors located in organs of the abdomen and thorax. The nodose ganglia of vagus nerves contain many types of receptors that modulate vagal activity. This protocol describes a method of local injections of neurochemicals into the nodose ganglia.

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located ...

Limbal Approach-Subretinal Injection of Viral Vectors for Gene Therapy in Mice Retinal Pigment Epithelium

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in retinopathies using non-viral and viral gene delivery to the retina and retinal pigment epithelium (RPE). Subretinal injection is the best approach to deliver viral vectors directly to RPE cells. Before the clinical trial of a gene therapy, it is inevitable to validate the efficacy of the therapy in animal models of various retinopathies. Thus, subretinal injection in mice becomes a fundamental technique for an ocular gene therapy. In this protocol, we provide the easy and replicable technique for subretinal injection of viral vectors to experimental mice. This technique is modified from the intravitreal injection, which is widely used technique in ophthalmology clinics. The representative results of RPE/choroid/scleral complex flat-mount will help to understand the efficacy of this technique and adjust the volume and titer of viral vectors for the extent of gene transduction.

Subretinal injection is a surgical technique for effective gene delivery to retinal pigment epithelium in the mouse eye. Here we describe an easy and replicable method for subretinal injection of viral vectors to retinal pigment epithelium in experimental mice.

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in ...

Conditions Affecting Social Space in Drosophila melanogaster

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a straightforward manner. As we previously demonstrated 1, in a two-dimensional chamber, we first force the flies to form a tight group, subsequently allowing them to take their preferred distance from each other. After the flies have settled, we measure the distance to the closest neighbor (or social space), processing a static picture with free online software (ImageJ). The analysis of the distance to the closest neighbor allows researchers to determine the effects of genetic and environmental factors on social interaction, while controlling for potential confounding factors. Diverse factors such as climbing ability, time of day, sex, and number of flies, can modify social spacing of flies. We thus propose a series of experimental controls to mitigate these confounding effects. This assay can be used for at least two purposes. First, researchers can determine how their favorite environmental shift (such as isolation, temperature, stress or toxins) will impact social spacing 1,2. Second, researchers can dissect the genetic and neural underpinnings of this basic form of social behavior 1,3. Specifically, we used it as a diagnostic tool to study the role of orthologous genes thought to be involved in social behavior in other organisms, such as candidate genes for autism in humans 4.

Conditions Affecting Social Space in Drosophila melanogaster

The effect of genes and environment on social space of Drosophila melanogaster can be quantified through a powerful but straightforward analytical paradigm. We show here different factors that can affect this social space, and thus need to be taken into consideration when designing experiments in this paradigm.

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a ...

A Simple Alternative to Stereotactic Injection for Brain Specific Knockdown of miRNA

MicroRNAs (miRNAs) are key regulators of gene expression. In the brain, vital processes like neurodevelopment and neuronal functions depend on the correct expression of microRNAs. Perturbation of microRNAs in the brain can be used to model neurodegenerative diseases by modulating neuronal cell death. Currently, stereotactic injection is used to deliver miRNA knockdown agents to specific location in the brain. Here, we discuss strategies to design antagomirs against miRNA with locked nucleotide modifications (LNA). Subsequently describe a method for brain specific delivery of antagomirs, uniformly across different regions of the brain. This method is simple and widely applicable since it overcomes the surgery, associated injury and limitation of local delivery in stereotactic injections. We prepared a complex of neurotropic, cell-penetrating peptide Rabies Virus Glycoprotein (RVG) with antagomir against miRNA-29 and injected through tail vein, to specifically deliver in the brain. The antagomir design incorporated features that allow specific targeting of the miRNA and formation of non-covalent complexes with the peptide. The knock-down of the miRNA in neuronal cells, resulted in apoptotic cell death and associated behavioural defects. Thus, the method can be used for acute models of neuro-degeneration through the perturbation of miRNAs.

MicroRNAs play crucial roles in the brain and are potential targets for modeling neuro-degeneration. However, perturbing miRNA levels is challenging due to the short length of miRNA and inaccessibility of the brain tissue. This video presents a method for antagomir design and brain specific delivery using a neuropeptide in mice.

MicroRNAs (miRNAs) are key regulators of gene expression.  In the brain, vital processes like neurodevelopment and neuronal functions depend on the ...

A New Approach that Eliminates Handling for Studying Aggression and the "Loser" Effect in Drosophila melanogaster

Aggressive behavior in Drosophila melanogaster is composed of the sequential expression of stereotypical behavioral patterns (for analysis see 1). This complex behavior is influenced by genetic, hormonal and environmental factors. As in many organisms, previous fighting experience influences the fighting strategy of flies and the outcome of later contests: losing a fight increases the probability of losing later contests, revealing &#34;loser&#34; effects that likely involve learning and memory 2-4. The learning and memory that accompanies expression of complex social behaviors like aggression, is sensitive to pre-test handling of animals 5,6. Many experimental procedures are used in different laboratories to study aggression 7-9, however, no routinely used protocol that excludes handling of flies is currently available. Here, we report a new behavioral apparatus that eliminates handling of flies, using instead their innate negative geotactic responses to move animals into or out of fighting chambers. In this protocol, small circular fight arenas containing a food cup are divided into two equal halves by a removable plastic slider prior to introduction of flies. Flies enter chambers from their home isolation vials via sliding chamber doors and geotaxis. Upon removal of plastic sliders, flies are free to interact. After specified time periods, flies are separated again by sliders for subsequent experimentation. All of this is done easily without handling of individual flies. This apparatus offers a novel approach to study aggression and the associated learning and memory, including the formation of &#34;loser&#34; effects in fly fights. In addition, this new general-purpose behavioral apparatus can be employed to study other social behaviors of flies and should, in general, be of interest for investigating experience-related changes in fundamental behavioral processes.

A New Approach that Eliminates Handling for Studying Aggression and the "Loser" Effect in Drosophila melanogaster

During fruit fly fights, the behavioral patterns observed, fight dynamics, and associated learning and memory are influenced by experimental conditions. The protocol presented here describes a novel procedure that entirely eliminates handling of flies during experiments. This improves fight dynamics and allows formation of strong &#34;loser&#34; effects.

Aggressive behavior in Drosophila melanogaster is composed of the sequential expression of stereotypical behavioral patterns (for analysis see 1). This ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, very few have been widely accepted. Here, we propose&#160;a new procedure&#160;adapted from the well-known ex vivo brain slice technique, which offers a closer in vivo-like scenario than in vitro preparations, for investigating the early events triggering cell degeneration, as observed in Alzheimer&#39;s disease (AD). This variation consists of simple and easily reproducible steps, which enable preservation of the anatomical cytoarchitecture of the selected brain region and its local functionality in a physiological milieu. Different anatomical areas can be obtained from the same brain, providing the opportunity to perform multiple experiments with the treatments in question in a site-, dose-, and time-dependent manner. Potential limitations&#160;which could affect the outcomes related to this methodology are related to the conservation of the tissue, i.e., the maintenance of its anatomical integrity during the slicing and incubation steps and the section thickness, which can influence the biochemical and immunohistochemical analysis. This approach can be employed for different purposes, such as exploring molecular mechanisms involved in physiological or pathological conditions, drug screening, or dose-response assays. Finally, this protocol could also reduce the number of animals employed in behavioral studies. The application reported here has been recently described and tested for the first time on ex vivo rat brain slices containing the basal forebrain (BF), which is one of the cerebral regions primarily affected in AD. Specifically, it has been demonstrated that the administration of a toxic peptide derived from the C-terminus of acetylcholinesterase (AChE) could prompt an AD-like profile, triggering, along the antero-posterior axis of the BF, a differential expression of proteins altered in AD, such as the alpha7 nicotinic receptor (&#945;7-nAChR), phosphorylated Tau (p-Tau), and amyloid beta (A&#946;).

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

We present a method which can&#160;provide further insights into the early events underlying neurodegeneration and based on the established ex-vivo brain technique, combining the advantages of in vivo and in vitro experiments. Moreover, it represents a unique opportunity for direct comparison of treated and untreated group in the same anatomical plane.

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, ...

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the underlying parenchyma. Previously published protocols recommend using straight needles that should be lowered to a maximum of 1-2 mm from the dural surface. The sudden drop in resistance once the dural membrane has been punctured makes it difficult to maintain the needle in a steady position. Our method, instead, employs a needle bent at the tip that can be stabilized against the occipital bone of the skull, thus preventing the syringe from penetrating into the tissue after perforation of the dural membrane. The procedure is straightforward, reproducible, and does not cause long-lasting discomfort in the operated animals. We describe the intracisternal injection strategy in the context of genetic fate mapping of vascular leptomeningeal cells. The same technique can, furthermore, be utilized to address a wide range of research questions, such as probing the role of leptomeninges in neurodevelopment and the spreading of bacterial meningitis, through genetic ablation of genes putatively implicated in these phenomena. Additionally, the procedure can be combined with an automatized infusion system for a constant delivery and used for tracking cerebrospinal fluid movement via injection of fluorescently labelled molecules.

We describe an intracisternal injection that employs a needle bent at the tip that can be stabilized to the skull, thus eliminating the risk of damage to the underlying parenchyma. The approach can be used for genetic fate mapping and manipulations of leptomeningeal cells and for tracking cerebrospinal fluid movement.

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the ...