This study was supported by grants from the Science and Technology Development Fund (FDCT) of Macao SAR (Ref. No. FDCT0058/2019/A1 and 0016/2019/AKP), Research Committee, University of Macau (MYRG2020-00183-ICMS and CPG2022-00023-ICMS), and National Natural Science Foundation of China (No. 81803398).

AB wild-type zebrafish and transgenic zebrafish lines elavl3:mCherry, ETvmat2:GFP, and mpo:EGFP were obtained from the Institute of Chinese Medical Sciences (ICMS). Ethical approval (UMARE-030-2017) for the animal experiments was granted by the Animal Research Ethics Committee, University of Macau, and the protocol follows the institutional animal care guidelines.
1. Zebrafish embryo and larval husbandry
<ol>
	<li>Generate zebrafish embryos (200-300 embryos per mating) by natural pairwise mating as previously reported28.</li>
	<li>Prepare egg water (E3) medium stock (60&#215;) by dissolving 34.8 g of NaCl, 1.6 g of KCl, 5.8 g of CaCl2&#183;2H2O, and 9.78 g of MgCl2&#183;6H2O in ddH2O to a final volume of 2 L and adjust pH to 7.2 using NaOH and HCl. To prepare working concentration of E3 medium (1&#215;), dilute the stock 60 times in ddH2O. Store at 28.5 &#176;C when not in use.</li>
	<li>Use a plastic transfer pipette to transfer the zebrafish embryos to a dish containing E3 medium. Raise the embryos at 28.5 &#176;C in an incubator and monitor their development and health. Remove dead embryos and replace unclean E3 medium with fresh medium daily.</li>
	<li>For zebrafish imaging experiments, use a plastic transfer pipette to collect 0-2 h post fertilization (hpf) embryos (around 200 eggs) in approximately 15 mL of 1&#215; E3 medium containing 0.003% N-phenylthiourea (PTU). 
	NOTE: PTU can inhibit the formation of melanophores during embryogenesis29. Treatment of embryos with PTU during embryogenesis can improve signal detection in microscopy or expression of fluorescence30.</li>
	<li>Maintain the zebrafish embryos under the above conditions until embryos reach the desired developmental larval stage.</li>
</ol>
2. Preparing for microinjection
<ol>
	<li>Prepare for the injection by pulling glass capillaries using a micropipette puller (see Table of Materials), following the five step protocol for glass capillary tube pulling (Table 1).</li>
	<li>Open the tip of needle (thin wall glass capillaries [3.5 in] with filament, OD 1.14 mm) to the appropriate size at an angle using forceps (Figure 2A).</li>
	<li>Fill the needle with 0.1 mL of mineral oil to ensure that there are no bubbles.</li>
	<li>Remove the screw cap of the steel needle of the microinjection apparatus. Align the glass needle hole with the steel needle and then tighten the screw cap. Mount the loaded needle microinjection apparatus in a micromanipulator (see Table of Materials).</li>
	<li>Adjust the position of the microinjection apparatus so that the micromanipulator can flexibly move the apparatus under the microscope. Discharge a certain volume of paraffin oil to make the steel needle enter the capillary glass tube.</li>
	<li>Drop the injection solution (such as 1&#215; PBS or 5 mg/mL LPS) on a glass slide sterilized with 70% ethanol and adjust the microinjection apparatus so that the tip is inserted into the liquid drop. Load ~2 &#181;L of injection solution into the needle.</li>
	<li>Set up the micromanipulator (fine adjustment settings of up, down, left, and right) so that the needle tip of the microinjection apparatus is in the same field of vision as the larvae on high magnification (Figure 2B).</li>
</ol>
3. Mounting zebrafish for microinjections
NOTE: Zebrafish brain development occurs within 3 dpf and matures at 5 dpf with a well-developed central nervous system31,32. Therefore, 5 dpf larvae are already suitable for studying LPS-mediated neuronal damage as well as behavioral and inflammatory responses.
<ol>
	<li>Inject the zebrafish larvae within approximately 30 min of reaching the stage of interest, to maintain the consistency of injection timing.</li>
	<li>To anesthetize the zebrafish larvae, combine tricaine with clean fish tank water (final concentration of 0.02% w/v tricaine).</li>
	<li>Melt a solution of 2% agarose in ddH2O using a microwave. Pour the molten agarose into a plastic dish. The 2% agarose-coated plastic dish can be stored at 4 &#176;C for up to 2 weeks.</li>
	<li>Use a plastic transfer pipette to transport anesthetized larvae to the center of the 2% agarose-coated plastic dish. Orient the mounted larvae with the brain side up for needle access.</li>
</ol>
4. Injecting the brain ventricle
<ol>
	<li>Adjust the magnification of the microscope so that the brain ventricular structure of zebrafish is clearly displayed in the field of vision (Figure 2B).</li>
	<li>Place the needle carefully above the brain tectum (Figure 2C).</li>
	<li>Puncture the skin of the zebrafish brain with the needle tip slowly using the micromanipulator.</li>
	<li>Press the foot pedal to eject 1 nL of 1x PBS or different concentrations of LPS (1.0, 2.5, and 5.0 mg/mL) (see Table of Materials). Successful ventricle injection images at which brain ventricle was injected with 1 nL of 1% Evans blue dye (diluted in PBS) are shown as an example (Figure 2D). Transfer the larvae to clean E3 medium immediately after the injection. After 24 h, collect the larvae for microscopic imaging, locomotive behavioral assay, and determination of other indicators.</li>
</ol>
5. Imaging
<ol>
	<li>Prepare 1.5% low-melt agarose solution in E3 medium and heat it in a microwave to form a clear liquid.</li>
	<li>Use a clean plastic transfer pipette to transport the larvae to a glass slide and remove as much water as possible.</li>
	<li>Use a plastic transfer pipette to add a drop of 1.5% low-melt agarose to the larvae. 
	NOTE: Cool the low-melt agarose in the pipette for a while before adding so as not to injure the larvae.</li>
	<li>Use a 1 mL syringe needle to orient the larvae. Wait until the agarose cools down and solidifies before starting imaging.</li>
	<li>Observe and photograph the neuron region in the whole brain of Tg(ETvmat2:EGFP) and Tg(elavl3:mCherry) zebrafish, as well as the recruitment of neutrophils in the Tg(mpo:EGFP) zebrafish brain under a fluorescence stereo microscope (see Table of Materials). 
	NOTE: Fluorescence microscope settings used for imaging are shown below. Tg(ETvmat2:EGFP) zebrafish brain (excitation wavelength: 450-490 nm, emission wavelength: 500-550 nm, visual magnification: 100x); Tg(elavl3:mCherry) zebrafish brain (excitation wavelength: 541-551 nm, emission wavelength: 590 nm, visual magnification: 100x); Tg(mpo:EFGP) zebrafish brain (excitation wavelength: 450-490 nm, emission wavelength: 500-550 nm, visual magnification: 63x).</li>
	<li>Measure and analyze the fluorescence intensity and length of the Ra neuron region in Tg(ETvmat2:EGFP) zebrafish and the fluorescence intensity of Tg(elavl3:mCherry) zebrafish brain neurons using ImageJ software. Manually count the neutrophils in the brain region of Tg(mpo:EFGP) zebrafish.</li>
</ol>
6. Determination of gene expression markers
<ol>
	<li>At 24 h after LPS brain ventricular injection, proceed with the determination of gene expression markers. To do so, anesthetize zebrafish larvae with 0.02% tricaine (as mentioned in step 3.2).</li>
	<li>Use two 1 mL syringes to separate the head portions of the larvae (40 larvae per group) without the eye and yolk sac regions (Figure 2E).</li>
	<li>Extract RNA from the larval head portions.
	<ol>
		<li>For RNA extraction, homogenize the head portion with 200 &#181;L of RNA extraction reagent (see Table of Materials) using a tissue grinder (see Table of Materials) at a rotation speed of 11,000 rpm for 5 s.</li>
		<li>Perform RNA extraction using the chloroform-isopropanol method. To do so, add 40 &#181;L chloroform, shake the tubes vigorously, and incubate them at room temperature for 10 min. Centrifuge the samples at 12,000 x g for 15 min at 4 &#176;C.</li>
		<li>Transfer the aqueous phase to a fresh tube (about 100 &#181;L) and add 100 &#181;L of isopropanol. Incubate for 10 min and then centrifuge at 12,000 x g for 10 min at 4 &#176;C. Remove the supernatant.</li>
		<li>Add 200 &#181;L of 75% ethanol to wash the RNA pellet. Vortex the samples briefly and then centrifuge at 7500 x g for 5 min at 4 &#176;C. Discard the supernatant and wash the RNA pellet again.</li>
		<li>Air dry the RNA pellet for 5-10 min, then add 30 &#181;L of RNase-free water to dissolve the RNA pellet. Check the purity (ratio of absorbance at 260 nm and 280 nm) and integrity of RNA using a microvolume spectrophotometer (see Table of Materials).</li>
	</ol>
	</li>
	<li>Synthesize cDNA from the RNA extracted in step 6.3, using reverse transcriptase with random primers (see Table of Materials).
	<ol>
		<li>Add 1 &#181;L of random primers, 1 &#181;L of dNTPs, 1 &#181;g of RNA, and distilled water (total volume to 12 &#181;L) to a nuclease-free tube. Heat the mixture to 65 &#176;C for 5 min and quick chill on ice.</li>
		<li>Add 1 &#181;L of 0.1 M DTT, 1 &#181;L of RNase inhibitor, 4 &#181;L of 5&#215; first-strand buffer, and 1 &#181;L of M-MLV reverse transcriptase (total volume: 20 &#181;L) to the tube. Mix gently and incubate the tube at 25 &#176;C for 2 min, followed by 42 &#176;C for 50 min. Inactivate the reaction by heating at 70 &#176;C for 15 min.</li>
	</ol>
	</li>
	<li>Perform real time PCR on a qPCR System (see Table of Materials) using a commercially available RT-qPCR kit (see Table of Materials) to determine the expression of the target genes(IL-1&#946;, IL-6, and TNF-&#945;). The reaction mixture (20 &#181;L) contained 10 &#181;L of SYBR green, 0.4 &#181;L of ROX reference dye, 0.8 &#181;L each of the forward and reverse primers, 6 &#181;L of ddH2O, and 2 &#181;L of template cDNA (1 &#181;g). Perform PCR amplification under the conditions: 95 &#176;C for 30 s, followed by 40 cycles of 95 &#176;C for 5 s and 60 &#176;C for 34 s, and with a dissociation stage of 95 &#176;C for 15 s, 60 &#176;C for 60 s, and 95 &#176;C for 15 s.</li>
	<li>Normalize the mRNA levels of the target genes to that of a housekeeping gene, elongation factor 1 &#945; (Ef1&#945;). Calculate the expression levels for each target gene by the 2&#8722;&#916;&#916;CT method33. The primer sequences of each gene are described in Table 2.</li>
	<li>For the determination of nitric oxide, homogenize the head portion (obtained in step 6.2) in 100 &#181;L of cold PBS using a tissue grinder. Centrifuge the resultant PBS and head portion suspension at 12,000 x g for 15 min at 4 &#176;C. Collect the lysates and perform nitrite concentration assay34 using a commercially available kit (see Table of Materials).</li>
</ol>
7. Zebrafish locomotive behavioral assay
<ol>
	<li>At 24 h after LPS brain ventricular injection, transfer the zebrafish larvae to the wells of a 96-well square microplate individually. Add 300 &#181;L of E3 medium to each well and then incubate the larvae for 4 h to acclimate to the testing plate.</li>
	<li>Transfer the larvae-loaded microplate to the zebrafish tracking box (see Table of Materials). Turn on the light source and then incubate the larvae in the testing box for 30 min to acclimate the environment.</li>
	<li>Monitor and record the zebrafish behavior using an automated video tracking system. Record 12 sessions (5 min each, total 1 h) for each zebrafish. Define the total distance (consists of inactive, small, and large distances) as the distance (in mm) that each fish moved during a 60 min tracking period.</li>
</ol>
8. Statistical analysis
<ol>
	<li>Perform statistical analyses using standard analysis software (see Table of Materials).</li>
	<li>Perform statistical analysis of differences between two groups using ordinary one-way ANOVA. Calculate Pearson&#39;s correlation coefficient to assess the strength of correlations. P &#60; 0.05 was considered significant in all analyses.</li>
</ol>

The authors declare no competing financial interest.

An increasing amount of epidemiological and experimental data implicate chronic bacterial and viral infections as possible risk factors for neurodegenerative diseases36. The infection triggers the activation of inflammatory processes and host immune responses37. Even if the response acts as a defense mechanism, overactivated inflammation is detrimental to neurogenesis, and the inflammatory environment does not allow for the survival of newborn neurons38</s...

Neuroinflammation has been described as a crucial anti-neurogenic factor involved in the pathogenesis of several neurodegenerative diseases of the central nervous system (CNS)1. Following pathological insults, neuroinflammation may result in various adverse consequences, including inhibition of neurogenesis and induction of neuronal cell death2,3. In the process underlying the response to inflammation induction, multiple inflammatory cytokines (such as TNF-&#945;, IL-1&#946;, and IL-6) are secreted into the extracellular space and act as crucial components in neuron death and the suppression of neurogenesis4,5,6.
Microinjection of inflammation mediators (such as IL-1&#946;, L-arginine, and endotoxins) into brain can cause neuronal cell reduction and neuroinflammation7,8,9. Lipopolysaccharide (LPS, Figure 1), a pathogenic endotoxin present in the cell wall of Gram-negative bacteria, can induce neuroinflammation, exacerbate neurodegeneration, and reduce neurogenesis in animals10. LPS injection directly into the CNS of the mouse brain increased levels of nitric oxide, pro-inflammatory cytokines, and other regulators11. Furthermore, stereotaxic injection of LPS into the local brain environment can induce excessive production of neurotoxic molecules, resulting in impaired neuronal function and subsequent development of neurodegenerative diseases10,12,13,14,15. In the neuroscience field, live and time-course microscopic observations of cellular and biological processes in living organisms are crucial for understanding the mechanisms underlying pathogenesis and pharmacological action16. However, live imaging of mouse models of neuroinflammation and neurotoxicity is fundamentally constrained by the limited optical penetration depth of microscopy, which precludes functional imaging and live observation of developmental processes17,18,19. Therefore, the development of alternative neuroinflammation models is of great interest to facilitate the study of pathological development, and the mechanism underlying neuroinflammation and neurodegeneration, by live imaging.
Zebrafish (Danio rerio) has emerged as a promising model to study neuroinflammation and neurodegeneration due its evolutionarily conserved innate immune system, optical transparency, large embryo clutch size, genetic tractability, and suitability for in vivo imaging19,20,21,22,23. Previous protocols have either directly injected LPS into the yolk and hindbrain ventricle of larval zebrafish without mechanistic assessment, or simply added LPS to fish water (culture medium) to induce a lethal systemic immune response24,25,26,27. Herein, we developed a protocol for microinjection of LPS into the brain ventricles, to trigger an innate immune response or neurotoxicity in the 5 days post fertilization (dpf) zebrafish larvae. This response is evidenced by neuronal cell loss, locomotory behavior deficit, increased nitrite oxide release, activation of inflammatory gene expression, and recruitment of neutrophils in the zebrafish brain at 24 h after injection.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Sigma-Aldrich</td>
			<td>A6361</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose, low gelling temperature</td>
			<td>Sigma-Aldrich</td>
			<td>A9414</td>
			<td></td>
		</tr>
		<tr>
			<td>Drummond Nanoject III Programmable Nanoliter Injector</td>
			<td>Drummond Scientific</td>
			<td>3-000-207</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence stereo microscopes</td>
			<td>Leica</td>
			<td>M205 FA</td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism software</td>
			<td>GraphPad Software</td>
			<td>Ver. 7.04</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipopolysaccharides from Escherichia coli O111:B4</td>
			<td>Sigma-Aldrich</td>
			<td>L3024</td>
			<td></td>
		</tr>
		<tr>
			<td>Manual micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>M3301</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Sigma-Aldrich</td>
			<td>M5904</td>
			<td></td>
		</tr>
		<tr>
			<td>Mx3005P qPCR system</td>
			<td>Agilent Technologies</td>
			<td>Mx3005P</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanovue plus spectrophotometer</td>
			<td>Biochrom</td>
			<td>80-2140-46</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitrite concentration assay kit</td>
			<td>Beyotime Biotechnology</td>
			<td>S0021M</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline</td>
			<td>Sigma-Aldrich</td>
			<td>P4417</td>
			<td></td>
		</tr>
		<tr>
			<td>Programmable Horizontal Pipette Puller</td>
			<td>World Precision Instruments</td>
			<td>PMP-102</td>
			<td></td>
		</tr>
		<tr>
			<td>PTU (N-Phenylthiourea)</td>
			<td>Sigma-Aldrich</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr>
			<td>Random primers</td>
			<td>Takara</td>
			<td>3802</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperScript II Reverse Transcriptase</td>
			<td>Invitrogen</td>
			<td>18064014</td>
			<td></td>
		</tr>
		<tr>
			<td>SYBR Premix Ex Taq II kit</td>
			<td>Accurate Biology</td>
			<td>AG11701</td>
			<td></td>
		</tr>
		<tr>
			<td>The 3rd Gen Tgrinder</td>
			<td>Tiangen</td>
			<td>OSE-Y30</td>
			<td></td>
		</tr>
		<tr>
			<td>Thin wall glass capillaries (4&#8221;) with filament, OD 1.5 mm</td>
			<td>World Precision Instruments</td>
			<td>TW150F-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine (3-amino benzoic acid ethyl ester)</td>
			<td>Sigma-Aldrich</td>
			<td>A-5040</td>
			<td></td>
		</tr>
		<tr>
			<td>TRNzol Universal reagent</td>
			<td>Tiangen</td>
			<td>DP424</td>
			<td></td>
		</tr>
		<tr>
			<td>Zebrafish tracking box</td>
			<td>ViewPoint Behavior Technology</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

brain ventricular microinjections of lipopolysaccharide into larval zebrafish to assess neuroinflammation and neurotoxicity

Neuroinflammation is a key player in various neurological disorders, including neurodegenerative diseases. Therefore, it is of great interest to research and develop alternative in vivo neuroinflammation models to understand the role of neuroinflammation in neurodegeneration. In this study, a larval zebrafish model of neuroinflammation mediated by ventricular microinjection of lipopolysaccharide (LPS) to induce an immune response and neurotoxicity was developed and validated. The transgenic zebrafish lines elavl3:mCherry, ETvmat2:GFP, and mpo:EGFP were used for real-time quantification of brain neuron viability by fluorescence live imaging integrated with fluorescence intensity analysis. The locomotor behavior of zebrafish larvae was recorded automatically using a video-tracking recorder. The content of nitric oxide (NO), and the mRNA expression levels of inflammatory cytokines including interleukin-6 (IL-6), interleukin-1&#946; (IL-1&#946;), and human tumor necrosis factor &#945; (TNF-&#945;) were investigated to assess the LPS-induced immune response in the larval zebrafish head. At 24 h after the brain ventricular injection of LPS, loss of neurons and locomotion deficiency were observed in zebrafish larvae. In addition, LPS-induced neuroinflammation increased NO release and the mRNA expression of IL-6, IL-1&#946;, and TNF-&#945; in the head of 6 days post fertilization (dpf) zebrafish larvae, and resulted in the recruitment of neutrophils in the zebrafish brain. In this study, injection of zebrafish with LPS at a concentration of 2.5-5 mg/mL at 5 dpf was determined as the optimum condition for this pharmacological neuroinflammation assay. This protocol presents a new, quick, and efficient methodology for brain ventricle microinjection of LPS to induce LPS-mediated neuroinflammation and neurotoxicity in a zebrafish larva, which is useful for studying neuroinflammation and could also be used as a high-throughput in vivo drug screening assay.

The workflow described here presents a new, quick, and efficient methodology for inducing LPS-mediated neuroinflammation and neurotoxicity in zebrafish larvae. In this described protocol, 5 dpf zebrafish were injected with LPS (Figure 1) into brain ventricles using a microinjector (Figure 2A-C). Successful injection into the brain ventricle site was verified using 1% Evans blue stain (Figure 2D). Th...

Watch this Scientific Journal Video about Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity at JoVE.com

Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity

This protocol demonstrates the microinjection of lipopolysaccharide into the brain ventricular region in a zebrafish larval model to study the resulting neuroinflammatory response and neurotoxicity.

Neuroinflammation is a key player in various neurological disorders, including neurodegenerative diseases. Therefore, it is of great interest to research ...

brain-ventricular-microinjections-lipopolysaccharide-into-larval

Research

JoVE Journal

Neuroscience

3.0K Views.  University of Macau, Avenida da Universidade. This protocol demonstrates the microinjection of lipopolysaccharide into the brain ventricular region in a zebrafish larval model to study the resulting neuroinflammatory response and neurotoxicity. 

Video: Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an injecting needle produces during its trajectory is a major concern especially in the mouse brain for not only the brain is small but also sometimes multiple injections are needed.  We show here a method to produce glass capillary needles with a 50-&mu;m lumen which significantly reduces the brain damage and allows a precise targeting into the rodent brain. This method allows a delivery of small volumes (from 20 to 100 nl), reduces bleeding risks, and minimizes passive diffusion of drugs into the brain parenchyma. By using different size of capillary glass tubes, or changing the needle lumen, several types of substances and cells can be injected. Microinjections with a glass capillary tube represent a significant improvement in injection techniques and deep brain targeting with minimal collateral damage in small rodents.

In this article, we show a method to make glass capillary needles with a 50-&mu;m lumen. This technique significantly reduces the brain damage, minimizes passive diffusion of drugs and allows a precise targeting into the rodent brain.

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an ...

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals are not viable in those cases where the modified gene is expressed or deleted in the whole organism. Moreover, a variety of compensatory mechanisms often make it difficult to interpret the results. The compensatory effects can be alleviated by either timing the gene expression or limiting the amount of transfected cells.
The method of postnatal non-ventricular microinjection and in vivo electroporation allows targeted delivery of genes, siRNA or dye molecules directly to a small region of interest in the newborn rodent brain. In contrast to conventional ventricular injection technique, this method allows transfection of non-migratory cell types. Animals transfected by means of the method described here can be used, for example, for two-photon in vivo imaging or in electrophysiological experiments on acute brain slices.

This protocol describes a non-viral method of delivery of genetic constructs to a certain area of living rodent brain. The method consists of plasmid preparation, micropipette fabrication, neonatal rat pup surgery, microinjection of the construct, and in vivo electroporation.

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Nerve Excitability Assessment in Chemotherapy-induced Neurotoxicity

Chemotherapy-induced neurotoxicity is a serious consequence of cancer treatment, which occurs with some of the most commonly used chemotherapies1,2. Chemotherapy-induced peripheral neuropathy produces symptoms of numbness and paraesthesia in the limbs and may progress to difficulties with fine motor skills and walking, leading to functional impairment. In addition to producing troubling symptoms, chemotherapy-induced neuropathy may limit treatment success leading to dose reduction or early cessation of treatment. Neuropathic symptoms may persist long-term, leaving permanent nerve damage in patients with an otherwise good prognosis3. As chemotherapy is utilised more often as a preventative measure, and survival rates increase, the importance of long-lasting and significant neurotoxicity will increase.
There are no established neuroprotective or treatment options and a lack of sensitive assessment methods. Appropriate assessment of neurotoxicity will be critical as a prognostic factor and as suitable endpoints for future trials of neuroprotective agents. Current methods to assess the severity of chemotherapy-induced neuropathy utilise clinician-based grading scales which have been demonstrated to lack sensitivity to change and inter-observer objectivity4. Conventional nerve conduction studies provide information about compound action potential amplitude and conduction velocity, which are relatively non-specific measures and do not provide insight into ion channel function or resting membrane potential. Accordingly, prior studies have demonstrated that conventional nerve conduction studies are not sensitive to early change in chemotherapy-induced neurotoxicity4-6. In comparison, nerve excitability studies utilize threshold tracking techniques which have been developed to enable assessment of ion channels, pumps and exchangers in vivo in large myelinated human axons7-9.
Nerve excitability techniques have been established as a tool to examine the development and severity of chemotherapy-induced neurotoxicity10-13. Comprising a number of excitability parameters, nerve excitability studies can be used to assess acute neurotoxicity arising immediately following infusion and the development of chronic, cumulative neurotoxicity. Nerve excitability techniques are feasible in the clinical setting, with each test requiring only 5 -10 minutes to complete. Nerve excitability equipment is readily commercially available, and a portable system has been devised so that patients can be tested in situ in the infusion centre setting. In addition, these techniques can be adapted for use in multiple chemotherapies.
In patients treated with the chemotherapy oxaliplatin, primarily utilised for colorectal cancer, nerve excitability techniques provide a method to identify patients at-risk for neurotoxicity prior to the onset of chronic neuropathy. Nerve excitability studies have revealed the development of an acute Na+ channelopathy in motor and sensory axons10-13. Importantly, patients who demonstrated changes in excitability in early treatment were subsequently more likely to develop moderate to severe neurotoxicity11. However, across treatment, striking longitudinal changes were identified only in sensory axons which were able to predict clinical neurological outcome in 80% of patients10. These changes demonstrated a different pattern to those seen acutely following oxaliplatin infusion, and most likely reflect the development of significant axonal damage and membrane potential change in sensory nerves which develops longitudinally during oxaliplatin treatment10. Significant abnormalities developed during early treatment, prior to any reduction in conventional measures of nerve function, suggesting that excitability parameters may provide a sensitive biomarker.

This abstract describes a novel method to assess the development of neurotoxicity in patients receiving chemotherapy treatment. While conventional assessment methods are limited in their ability to detect early changes in nerve function, nerve excitability techniques provide early identification of patients at risk of severe neurotoxicity and insight into pathophysiology.

Chemotherapy-induced neurotoxicity is a serious consequence of cancer treatment, which occurs with some of the most commonly used chemotherapies1,2. ...

Manual Drainage of the Zebrafish Embryonic Brain Ventricles

Cerebrospinal fluid (CSF) is a protein rich fluid contained within the brain ventricles. It is present during early vertebrate embryonic development and persists throughout life. Adult CSF is thought to cushion the brain, remove waste, and carry secreted molecules1,2. In the adult and older embryo, the majority of CSF is made by the choroid plexus, a series of highly vascularized secretory regions located adjacent to the brain ventricles3-5. In zebrafish, the choroid plexus is fully formed at 144 hours post fertilization (hpf)6. Prior to this, in both zebrafish and other vertebrate embryos including mouse, a significant amount of embryonic CSF (eCSF) is present . These data and studies in chick suggest that the neuroepithelium is secretory early in development and may be the major source of eCSF prior to choroid plexus development7.
eCSF contains about three times more protein than adult CSF, suggesting that it may have an important role during development8,9. Studies in chick and mouse demonstrate that secreted factors in the eCSF, fluid pressure, or a combination of these, are important for neurogenesis, gene expression, cell proliferation, and cell survival in the neuroepithelium10-20. Proteomic analyses of human, rat, mouse, and chick eCSF have identified many proteins that may be necessary for CSF function. These include extracellular matrix components, apolipoproteins, osmotic pressure regulating proteins, and proteins involved in cell death and proliferation21-24. However, the complex functions of the eCSF are largely unknown.
We have developed a method for removing eCSF from zebrafish brain ventricles, thus allowing for identification of eCSF components and for analysis of the eCSF requirement during development. Although more eCSF can be collected from other vertebrate systems with larger embryos, eCSF can be collected from the earliest stages of zebrafish development, and under genetic or environmental conditions that lead to abnormal brain ventricle volume or morphology. Removal and collection of eCSF allows for mass spectrometric analysis, investigation of eCSF function, and reintroduction of select factors into the ventricles to assay their function. Thus the accessibility of the early zebrafish embryo allows for detailed analysis of eCSF function during development.

We present a method to collect cerebrospinal fluid (CSF) and to create a system which lacks CSF within the embryonic zebrafish brain ventricular system. This allows for further examination of CSF composition and its requirement during embryonic brain development.

Cerebrospinal fluid  (CSF) is a protein rich fluid contained within the brain ventricles. It is  present during early vertebrate embryonic development and ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Selective Depletion of Microglia from Cerebellar Granule Cell Cultures Using L-leucine Methyl Ester

Microglia, the resident immunocompetent cells of the CNS, play multifaceted roles in modulating and controlling neuronal function, as well as mediating innate immunity. Primary rodent cell culture models have greatly advanced our understanding of neuronal-glial interactions, but only recently have methods to specifically eliminate microglia from mixed cultures been utilized. One such technique &#8211; described here &#8211; is the use of L-leucine methyl ester, a lysomotropic agent that is internalized by macrophages and microglia, wherein it causes lysosomal disruption and subsequent apoptosis13,14. Experiments using L-leucine methyl ester have the power to identify the contribution of microglia to the surrounding cellular environment under diverse culture conditions. Using a protocol optimized in our laboratory, we describe how to eliminate microglia from P5 rodent cerebellar granule cell culture. This approach allows one to assess the relative impact of microglia on experimental data, as well as determine whether microglia are playing a neuroprotective or neurotoxic role in culture models of neurological conditions, such as stroke, Alzheimer&#8217;s or Parkinson&#8217;s disease.

Microglia can influence neurons and other glia in culture by various non-cell autonomous mechanisms. Here, we present a protocol to selectively deplete microglia from primary neuronal cultures. This method has the potential to elucidate the role of microglial-neuronal interactions, with implications for neurodegenerative conditions where neuroinflammation is a hallmark feature.

Microglia, the resident immunocompetent cells of the CNS, play multifaceted roles in modulating and controlling neuronal function, as well as mediating ...

Microglia as a Surrogate Biosensor to Determine Nanoparticle Neurotoxicity

Nanoparticles found in air pollutants can alter neurotransmitter profiles, increase neuroinflammation, and alter brain function. Therefore, the assay described here will aid in elucidating the role of microglia in neuroinflammation and neurodegenerative diseases. The use of microglia, resident immune cells of the brain, as a surrogate biosensor provides novel insight into how inflammatory responses mediate neuronal insults. Here, we utilize an immortalized murine microglial cell line, designated BV2, and describe a method for nanoparticle exposure using silver nanoparticles (AgNPs) as a standard. We describe how to expose microglia to nanoparticles, how to remove nanoparticles from supernatant, and how to use supernatant from activated microglia to determine toxicity, using hypothalamic cell survival as a measure. Following AgNP exposure, BV2 microglial activation was validated using a tumor necrosis factor alpha (TNF-&#945;) enzyme linked immunosorbent assay (ELISA). The supernatant was filtered to remove the AgNP and to allow cytokines and other secreted factors to remain in the conditioned media. Hypothalamic cells were then exposed to supernatant from AgNP activated microglia and survival of neurons was determined using a resazurin-based fluorescent assay. This technique is useful for utilizing microglia as a surrogate biomarker of neuroinflammation and determining the effect of neuroinflammation on other cell types.

Microglia (immune cells of the brain), are used as a surrogate biosensor to determine how nanoparticles influence neurotoxicity. We describe a series of experiments designed to assay microglial response to nanoparticles and exposure of hypothalamic neurons to supernatant from activated microglia to determine neurotoxicity.

Nanoparticles found in air pollutants can alter neurotransmitter profiles, increase neuroinflammation, and alter brain function. Therefore, the assay ...

Personalized Needles for Microinjections in the Rodent Brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity

Summary

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Chapters in this video

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nerve Excitability Assessment in Chemotherapy-induced Neurotoxicity

Manual Drainage of the Zebrafish Embryonic Brain Ventricles

Forebrain Electrophysiological Recording in Larval Zebrafish

Spinal Cord Transection in the Larval Zebrafish

Selective Depletion of Microglia from Cerebellar Granule Cell Cultures Using L-leucine Methyl Ester

Microglia as a Surrogate Biosensor to Determine Nanoparticle Neurotoxicity

Personalized Needles for Microinjections in the Rodent Brain