This work is supported by the BBSRC, EPSRC, MRC and Wellcome Trust. The C.B. laboratory is part of the Wellcome Trust Centre for Cell-Matrix research, University of Manchester, which is supported by core funding from the Wellcome Trust (grant number 088785/Z/09/Z). The authors wish to acknowledge the funding provided by the Biotechnology and Biological Sciences Research Council (BBSRC) to C.M., K.J. (BB/M020630/1) and P.A. (BB/P000681/1) and by the Engineering and Physical Sciences Research Council (EPSRC) and Medical Research Council (MRC) Centre for Doctoral Training in Regenerative Medicine to A.K. (EP/L014904/1). The authors thank Alv&#233;ole for their correspondence and their after-sales support team. The authors thank Peter March and Roger Meadows from the Bioimaging Facility, University of Manchester for their help with the microscopy. The Bioimaging Facility microscopes used in this study were purchased with grants from BBSRC, Wellcome Trust and the University of Manchester Strategic Fund.

The authors have nothing to disclose.

LIMAP (PRIMO) micro-patterning advantages and comparison with microcontact printing 
While microcontact printing is possibly the most commonly used micro-patterning technique in the biological field39, there seems to be an increasing number of researchers using LIMAP technology40,41,42,43,44. Here, we presented a protocol using PRIMO, a commercially available system for LIMAP. Below we briefly discuss potential advantages and limitations of microcontact printing and LIMAP photo-patterning.
Microcontact printing requires lithographed masters produced by spin-coating a photo-mask (generally SU-8) onto a glass or silicon wafer, which is then laser-etched with the desired micro-features. These masters are used as templates to create a PDMS stamp45. The stamp is incubated with a chosen protein that adsorbs to it, and is then transferred (stamped) onto the cell culture dish. The process of adsorption of the protein to the PDMS stamp is dependent on protein concentration, buffer and incubation time. These parameters need to be tested beforehand for optimal results46.
Masters can be used in a substantial number of experiments, lasting for months or even years, if correctly preserved. However, a limiting factor of this technology is the necessity to re-design new lithographed masters for every desired modification. Changes in experimental designs may result in time-consuming production of new masters (up to several weeks) thus delaying experiments. In comparison, LIMAP photopatterning does not require a physical master; it uses software-generated pattern templates that can be used to flexibly adapt desired geometries of micro-patterns to changing research questions. LIMAP can be also used to generate protein gradients within the same micro-pattern (Figure 8), which is harder to obtain in a reproducible manner using microcontact printing47.
Furthermore, the micro-pattern resolution achieved with LIMAP, in our case, is 2 &#181;m (Figure 6B).
Approaching this resolution increased intra- and inter- pattern variability. Generating patterns around or above 10 &#181;m width was highly reproducible (Figure 6G,H). On the contrary, with microcontact printing it is difficult to consistently obtain resolutions below 10 &#181;m and it is common to find artefacts when stamping small features (data not shown).
We have shown that LIMAP can be used to micro-pattern multiple proteins (Figure 9) within the same micro-well, allowing further levels of complexity to be added to experiments. Although this could be achieved with microcontact printing, aligning different proteins with high level of precision can be technically rather demanding. Whilst patterning multiple proteins using LIMAP seems straight forward, it is important to mention that cross-binding of proteins through sequential coating procedures can be reduced through blocking reagents but not entirely eliminated (Figure 9).
Regarding the cost of one or the other technique, LIMAP as described here requires the purchase of micro-patterning equipment (PRIMO) that can be installed on different fluorescence microscopes and requires a motorized stage. Although this investment is initially cost intensive, there are no additional purchases other than consumable items (stencils, PEG and PLPP) on the long run associated with LIMAP. Alternatively, the PDMS stencils can also be produced in the lab by the own experimenter following published protocols18,32. The largest costs for microcontact printing may be associated with the production of new masters, which can become substantial if experiments require new patterns.
One drawback of LIMAP is the relatively low throughput approach of this technique. Microcontact printing can produce a large number of micro-patterns quickly and efficiently in a simultaneous stamping step, compared to the required sequential laser micro-patterning with LIMAP. For example, it is possible to produce 6 stamped glass coverslips in about 2 h with microcontact printing using PDMS stamps (excluding stamp preparation); patterning a similar area (6-well dish) with LIMAP would take around 4 h, excluding the procedure of surface passivation (considering the pattern template configuration described in step 5.12 and see Figure 5B).
Another rate limiting factor of LIMAP technology is the long illumination time required for patterning large areas (30 s per design unit with a 7.5 mW/mm2 laser). In these cases, microcontact printing might be a preferred option. A newly available photo-initiator (PLPP gel, Table of Materials) should considerably reduce the time taken for the patterning, allowing the generation of hundreds of micro-patterns in large areas (up to 8 mm2) in just a few minutes.
Another important factor to take into account when micro-patterning surfaces for cell culture is the reproducibility of the micro-patterns among different experimental repeats, in comparison to the variability obtained with microcontact printing. For example, the graphs shown in Figure 7B,D are representative data of three independent experimental repeats with very similar results (data not shown). Based on our experience and previous publications, this level of reproducibility is difficult to achieve with microcontact printing48,49,50,51,52.
In contrast to other photo-patterning techniques that require either dedicated chemistry to engineer photosensitive materials or the use of photo-sensitizers, which are generally not very biocompatible3, the photosensitive component of LIMAP (PLPP) is biocompatible and well-tolerated by cells21; in our hands we have not experienced any cytotoxicity across a variety of cells, including CAD, DRG neurons (Figure 10), fibroblasts, epithelial cells, and melanoma cells (data not shown). Another advantage of LIMAP using PRIMO compared to other photo-patterning techniques is that no photomask is required. Similar to microcontact printing, new photomasks would need to be designed and generated for every desired pattern.
All the limitations mentioned above for microcontact printing, refer to the manual approach of the technique. However, it is possible to enhance the throughput and the reproducibility of microcontact printing using an automated device with stamp load and pressure control53.
Key steps of the protocol and problems solving for LIMAP using PRIMO 
One of the most common problems found during this protocol is having high levels of background fluorescence within the micro-patterns. This can be due to the drying out of micro-wells which often occurs due to their small volume. When this occurs, PBS crystals often appear surrounding the ECM patterns (Figure 11A).
Insufficient or inefficient washing steps after protein incubation can also result in high levels of background fluorescence. This can be observed particularly upon using protein concentrations of 10 &#181;g/mL (Figure 11B) or higher. The excess of protein in the background can be reduced by including additional washing steps with PBS.
The presence of protein background needs to be measured and characterized in each experiment, calculating the background fluorescence intensity (Figure 6E) and subtracting it from the micro-patterns intensity (Figure 6F-H and Figure 7B,D). High protein background may have an impact in the attachment and sprouting of CAD cells, compromising the interpretation of results.
Having gaps between design units is a common problem when users have limited experience (Figure 11B), which occurs as a result of insufficient overlap between patterns. Two parameters in the Leonardo software can be adjusted to overcome this: 1) a negative spacing between columns may be required, depending on the design of the pattern (step 5.7 and see Figure 5B,C). Alternatively, 2) use the gradient option in the Expert menu to stitch the columns. A quick test to determine the optimal spacing parameters can be performed using UV adhesive (Table of Materials). A small drop of this adhesive is applied to a glass slide, which is then covered with a glass coverslip, making a film. The embedded UV adhesive is photopatterned with the pattern template of interest using a low laser dose (30 mJ/mm2). The UV- exposed regions of the embedded adhesive will be cured, becoming visible under bright-field microscopy. The test results are visualized to evaluate the obtained spacing within the pattern. In our neuronal experiments, a gap between stripes may adversely affect cell behavior, producing variations in growth dynamics (either reduced speed or abandonment of the path).
In the latest update of Leonardo software (at the time of publication, Leonardo 4.11), it is possible to upload previously designed bigger pattern templates that cover a much larger area (up to 8 mm2 using the 20X objective) of the micro-well surface compared to the current 0.1 mm2 per design unit, eliminating the need to stitch together the smaller design units. Undefined edges can result from a lack of laser focus adjustment during pattern generation (Figure 11C). It is therefore critical to calibrate the laser and perform reference pattern steps (see step 4) prior to patterning. Poorly defined stripes result in variations in stripe width, making the correlation between axon growth dynamics and stripe width difficult. Axons also tend to abandon stripes that have diffused edges. Additionally, variability in edges can also be found when printing stripes of 10-20 &#181;m width or higher, resulting in a higher protein content at the edges compared to the central regions of the pattern (Figure 6B,D). This edge effect is produced by a non-homogeneous diffusion of the photo-initiator during the photopatterning process. The photoscission reaction is oxygen dependent, which diffuses more at the edges. This edge effect can be minimized homogenizing the photo-initiator with a pipette in the micro-well during the photopatterning process. Furthermore, a new commercialized photo-initiator (PLPP gel), can also reduce the edge effect (PRIMO system support team, personal communication).
Micro-printing of more than one protein can result in cross-binding (Figure 9A-D). This can be minimized by increasing the blocking efficiency that is used to occupy unspecific binding sites between the incubation steps for the two different proteins. Cross-binding of proteins can perturb reproducibility of experimental outcomes and may lead to misinterpretation of data, since it is difficult to determine the contribution of each protein to axon growth dynamics and to other cell behaviors.
Conclusion 
We hope that the provided protocol using LIMAP facilitates the generation of protein micro-patterns through the use of the PRIMO system. Whilst our protocol focuses on how to reliably produce micro-patterns in 2D glass surfaces, others have shown that it is possible to use LIMAP for micro-patterning of soft substrates54, and microstructured surfaces for 3D cultures42. These micro-patterns can be a versatile tool to study cellular responses to changes in their micro-environment.

In recent years, the biological sciences have increasingly made use of the advances provided by material sciences. One prominent example is the micro-patterning of substrates, which can be used to study cellular responses such as cell proliferation1,2, differentiation3,4,5,6, cell migration7,8,9 and pathfinding10,11. There are a number of techniques available that enable the micro-patterning of substrates, such as multiphoton excited photochemistry12, AFM dip-pen nanolithography13, pin and inkjet direct printing14, electron beam lithography15 or microfluidics16. However, two techniques that are widely used in the biological field are microcontact printing17,18,19 or laser-assisted patterning3 (Figure 1). Laser-assisted patterning is considered to deliver more reliable results in terms of protein and PEG stability and cell confinement on the patterns, compared to microcontact printing20. A more novel approach for micro-patterning described here is the use of Light-Induced Molecular Adsorption of Proteins21 (LIMAP, Figure 1D) using a commercially available system (PRIMO, Table of Materials). Each of the methods has advantages and limitations that are briefly described below.Microcontact printing uses PDMS molds (stamps) with desired micro-features that are generated from lithographed masters. The stamps are incubated with a chosen protein which is then transferred (stamped) onto the cell culture substrate18 (Figure 1A). Laser-assisted patterning uses UV light to cleave an anti-fouling film22,23,24,25, exposing regions that can subsequently be coated with the protein of interest (Figure 1B). While the resolution achieved with photo-patterning approaches is in the micron range25,26, most of these techniques require a photo-mask, either in contact with the sample, or situated in the object plane of the microscope objective23,27,28. The requirements for masks in both microcontact printing and photo-patterning can be a limitation; specific masks are required for every geometric pattern and size, which can be expensive and time consuming to generate. In contrast to these techniques, LIMAP does not require a mask (Figure 1D). Using the PRIMO system for LIMAP can be cost intensive at the beginning because it requires purchasing of equipment. However, open-source software is used to design patterns of any desired geometry, giving much more freedom and allowing more complex experiments including the use of protein concentration gradients.The PRIMO laser is controlled and directed by a digitally-controlled micromirror device (DMD) to create patterns in any number of user-defined geometries. LIMAP requires the culture surface to be coated with molecules that prevent cell attachment. Polyethylene glycol (PEG) is most commonly used as such an &#34;antifouling&#34; reagent; it forms a dense anti-adhesive film on the glass or plastic surface29. Subsequently, a photo-initiator is added that allows the PEG film to be removed with high precision through a photoscission mechanism30 by local exposure to UV light under the control of the DMD. These PEG-free regions can be coated with proteins that adsorb to the laser-etched surface, generating a micro-pattern. By varying the laser power, different quantities of PEG can be removed from the surface allowing the user to generate protein gradients. PEG removal and the coating procedure can be repeated to create patterns with two or more distinct proteins in the same micro-well21.The generated micro-patterns provide adhesive surfaces for cells, allowing the study of cell behavior. In our studies, we use micro-patterning to study neurite or axon pathfinding of a neuronal cell line (CAD (Cathecholaminergic-a differentiated) cells31) or primary rat dorsal-root ganglion (DRG) neurons, respectively. Here, we outline a step-by-step protocol for LIMAP (Figure 2) using the commercially available PRIMO system and accompanying Leonardo software. We demonstrate how it can be used for the generation of patterns with defined geometries and multiple proteins, which we use to study axonal pathfinding. We discuss common issues that can arise and how these can be avoided.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa 488 protein labeling kit</td>
			<td>Invitrogen</td>
			<td>A10235</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Alexa 647 protein labeling kit</td>
			<td>Invitrogen</td>
			<td>A20173</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>CAD cells</td>
			<td>ECACC</td>
			<td>8100805</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Conjugated fibrinogen-488</td>
			<td>Molecular Probes</td>
			<td>F13191</td>
			<td>Working concentration: 10 &#956;g/ml</td>
		</tr>
		<tr>
			<td>DMEM culture medium</td>
			<td>Gibco</td>
			<td>11320033</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Epifluorescence Microscope**</td>
			<td>Nikon</td>
			<td>Eclipse Ti inverted</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma</td>
			<td>F4759</td>
			<td>Working concentration: 10 &#956;g/ml 
			(after labelling with Alexa 488 protein labeling kit, see above) 
			(diluted in PBS)</td>
		</tr>
		<tr>
			<td>Fiji-Image J</td>
			<td><a href="http://www.imagej.nih.gov" target="_blank">www.imagej.nih.gov</a></td>
			<td>Version 2.0.0-rc-54/1.51f</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Fluorescent highlighter</td>
			<td>Stabilo</td>
			<td>Stabilo Boss Original</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Gibco</td>
			<td>15630080</td>
			<td>Working concentration: 1M</td>
		</tr>
		<tr>
			<td>Inkscape software</td>
			<td>Inkscape</td>
			<td>Check last update</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Laminin-red fluorescent rhodamine</td>
			<td>Cytoskeleton, Inc.</td>
			<td>LMN01</td>
			<td>Working concentration: 10 &#956;g/ml 
			(diluted in PBS)</td>
		</tr>
		<tr>
			<td>Leonardo software</td>
			<td>Alv&#233;ole</td>
			<td>version 4.11</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Sigma</td>
			<td>G7513</td>
			<td>Working concentration: 1%</td>
		</tr>
		<tr>
			<td>Micro-manager software</td>
			<td>Open imaging</td>
			<td>Check last update</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Motorized x/y stage</td>
			<td>PRIOR Scientific</td>
			<td>Proscan II</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>NIS Elements Software</td>
			<td>Nikon</td>
			<td>NIS Elements AR 4.60.00 64-bit (With Nikon jobs)</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>PBS (without Ca2+, Mg2+)</td>
			<td>Sigma</td>
			<td>D8537</td>
			<td>Working concentration: 1X</td>
		</tr>
		<tr>
			<td>PDMS Stencils</td>
			<td>Alv&#233;ole</td>
			<td>visit <a href="http://www.alveolelab.com" target="_blank">www.alveolelab.com</a></td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>PEG-SVA</td>
			<td>Laysan bio, Inc.</td>
			<td>MPEG-SVA-5000-1g</td>
			<td>Working concentration: 50 mg/ml</td>
		</tr>
		<tr>
			<td>Phalloidin 405</td>
			<td>Abcam</td>
			<td>ab176752</td>
			<td>Working concentration: 1:1000</td>
		</tr>
		<tr>
			<td>Photo-initiator (PLPP)</td>
			<td>Alv&#233;ole</td>
			<td>Classic PLPP</td>
			<td>Working concentration: 14.5 mg/ml</td>
		</tr>
		<tr>
			<td>Photo-initiator (PLPP gel)</td>
			<td>Alv&#233;ole</td>
			<td>PLPP gel</td>
			<td>Working concentration: 4.76% diluted in ethanol</td>
		</tr>
		<tr>
			<td>Plasma cleaner</td>
			<td>Harrick Plasma</td>
			<td>PDC-32G (115V) | PDC-32G-2 (230V)</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>PLL-PEG</td>
			<td>SuSoS (also distributed by Alv&#233;ole)</td>
			<td><a href="http://www.alveolelab.com" target="_blank">www.alveolelab.com</a></td>
			<td>Working concentration: 0.1 mg/ml (diluted in PBS)</td>
		</tr>
		<tr>
			<td>Poly-L-Lysine</td>
			<td>Sigma</td>
			<td><a href="https://www.sigmaaldrich.com/ProductLookup.html?ProdNo=P4707&amp;Brand=SIGMA">P4707</a></td>
			<td>Working concentration: 0.01%</td>
		</tr>
		<tr>
			<td>Primo equipment</td>
			<td>Alv&#233;ole</td>
			<td><a href="http://www.alveolelab.com" target="_blank">www.alveolelab.com</a></td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>Pen/Strep</td>
			<td>Thermo Fisher</td>
			<td><a href="https://www.thermofisher.com/order/catalog/product/15140122">15140122</a></td>
			<td>Working concentration: 1%</td>
		</tr>
		<tr>
			<td>Tubulin anti-alpha antibody</td>
			<td>Abcam</td>
			<td>DM1A</td>
			<td>Working concentration: 1:1000 
			CAD cells</td>
		</tr>
		<tr>
			<td>Tubulin anti-beta 3 antibody</td>
			<td>Sigma</td>
			<td>T8660</td>
			<td>Working concentration: 1:500 
			DRG neurons</td>
		</tr>
		<tr>
			<td>UV adhesive</td>
			<td>Norland Products</td>
			<td>NOA81</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>1 well glass bottom dish</td>
			<td>Cellvis</td>
			<td>D35-20-1.5-N</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>6 well glass bottom dish</td>
			<td>Cellvis</td>
			<td>P06-20-1.5-N</td>
			<td>Working concentration: N.A.</td>
		</tr>
		<tr>
			<td>20x objective**</td>
			<td>Nikon</td>
			<td>no phase ring (check updated catalogue)</td>
			<td>Working concentration: N.A. 
			**Epifluorescence microscope: images were acquired and patterns were generated on an Eclipse Ti inverted microscope (Nikon), coupled to PRIMO micro-patterning equipment (Alv&#233;ole), using a 20x objective (0.75 S Plan Fluor (nophasering, Nikon). Nikon specific filter sets for GFP, mCherry and Cy5 were used and fluorescent light source was LED (Lumencor) although other fluorescence sources and filter sets can be used. The microscope has an automated x/y stage (PRIOR Scientific) for the printing of multi-field patterning and Nikon Perfect Focus to prevent focus drift. The images were collected using a Retiga R6 (Q-Imaging) camera.</td>
		</tr>
	</tbody>
</table>

light-induced molecular adsorption of proteins using the primo system for micro-patterning to study cell responses to extracellular matrix proteins

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the cells themselves. Here, we present the method of Light-Induced Molecular Adsorption of Proteins (LIMAP) using the PRIMO system as a patterning technique to produce micro-patterned extracellular matrix (ECM) substrates using a single or combination of proteins. The method enables printing of ECM patterns in micron resolution with excellent reproducibility. We provide a step-by-step protocol and demonstrate how this can be applied to study the processes of neuronal pathfinding. LIMAP has significant advantages over existing micro-printing methods in terms of the ease of patterning more than one component and the ability to generate a pattern with any geometry or gradient. The protocol can easily be adapted to study the contribution of almost any chemical component towards cell fate and cell behavior. Finally, we discuss common issues that can arise and how these can be avoided.

Watch this Scientific Journal Video about Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins at JoVE.co

Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins

Our overall aim is to understand how cells sense extracellular cues that lead to directed axonal growth. Here, we describe the methodology of Light-Induced Molecular Adsorption of Proteins, used to produce defined micro-patterns of extracellular matrix components in order to study specific events that govern axon outgrowth and pathfinding.

Cells sense a variety of extracellular cues, including the composition and geometry of the extracellular matrix, which is synthesized and remodeled by the ...

light-induced-molecular-adsorption-proteins-using-primo-system-for

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Neuroscience

11.8K Views.  The University of Manchester. Our overall aim is to understand how cells sense extracellular cues that lead to directed axonal growth. Here, we describe the methodology of Light-Induced Molecular Adsorption of Proteins, used to produce defined micro-patterns of extracellular matrix components in order to study specific events that govern axon outgrowth and pathfinding.

Video: Light-Induced Molecular Adsorption of Proteins Using the PRIMO System for Micro-Patterning to Study Cell Responses to Extracellular Matrix Proteins

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

Isolation of CA1 Nuclear Enriched Fractions from Hippocampal Slices to Study Activity-dependent Nuclear Import of Synapto-nuclear Messenger Proteins

Studying activity dependent protein expression, subcellular translocation, or phosphorylation is essential to understand the underlying cellular mechanisms of synaptic plasticity. Long-term potentiation (LTP) and long-term depression (LTD) induced in acute hippocampal slices are widely accepted as cellular models of learning and memory. There are numerous studies that use live cell imaging or immunohistochemistry approaches to visualize activity dependent protein dynamics. However these methods rely on the suitability of antibodies for immunocytochemistry or overexpression of fluorescence-tagged proteins in single neurons. Immunoblotting of proteins is an alternative method providing independent confirmation of the findings.&#160;The first limiting factor in preparation of subcellular fractions from individual tetanized hippocampal slices is the low amount of material. Second, the handling procedure is crucial because even very short and minor manipulations of living slices might induce activation of certain signaling cascades. Here we describe an optimized workflow in order to obtain sufficient quantity of nuclear enriched fraction of sufficient purity from the CA1 region of acute hippocampal slices from rat brain. As a representative example we show that the ERK1/2 phosphorylated form of the synapto-nuclear protein messenger Jacob actively translocates to the nucleus upon induction of LTP and can be detected in a nuclear enriched fraction from CA1 neurons.

We provide a detailed protocol for induction of long-term potentiation in the CA1 region of the hippocampus and the subsequent isolation of nuclear enriched fractions from the tetanized area of the slice. This approach can be used to determine activity dependent nuclear protein import in cellular models of learning and memory.

Studying activity dependent protein expression, subcellular translocation, or phosphorylation is essential to understand the underlying cellular ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, their distinct dendritic morphologies, their specific patterns of axonal connectivity, and their selective firing responses. The molecular and cellular mechanisms responsible for these aspects of differentiation during development are still poorly understood.
Here, we describe combined protocols for labeling and characterizing the structural connectivity and excitability of cortical neurons. Modification of the in utero electroporation (IUE) protocol allows the labeling of a sparse population of neurons. This, in turn, enables the identification and tracking of the dendrites and axons of individual neurons, the precise characterization of the laminar location of axonal projections, and morphometric analysis. IUE can also be used to investigate changes in the excitability of wild-type (WT) or genetically modified neurons by combining it with whole-cell recording from acute slices of electroporated brains. These two techniques contribute to a better understanding of the coupling of structural and functional connectivity and of the molecular mechanisms controlling neuronal diversity during development. These developmental processes have important implications on axonal wiring, the functional diversity of neurons, and the biology of cognitive disorders.

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

This manuscript provides protocols that use in utero electroporation (IUE) to describe the structural connectivity of neurons at the single-cell level and the excitability of fluorescently labeled neurons. Histology is used to characterize dendritic and axonal projections. Whole-cell recording in acute slices is used to investigate excitability.

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

In Vivo Microdialysis Method to Collect Large Extracellular Proteins from Brain Interstitial Fluid with High-molecular Weight Cut-off Probes

In vivo microdialysis is a powerful technique to collect ISF from awake, freely-behaving animals based on a dialysis principle. While microdialysis is an established method that measures relatively small molecules including amino acids or neurotransmitters, it has been recently used to also assess dynamics of larger molecules in ISF using probes with high molecular weight cut off membranes. Upon using such probes, microdialysis has to be run in a push-pull mode to avoid pressure accumulated inside of the probes. This article provides step-by-step protocols including stereotaxic surgery and how to set up microdialysis lines to collect proteins from ISF. During microdialysis, drugs can be administered either systemically or by direct infusion into ISF. Reverse microdialysis is a technique to directly infuse compounds into ISF. Inclusion of drugs in the microdialysis perfusion buffer allows them to diffuse into ISF through the probes while simultaneously collecting ISF. By measuring tau protein as an example, the author shows how its levels are altered upon stimulating neuronal activity by reverse microdialysis of picrotoxin. Advantages and limitations of microdialysis are described along with the extended application by combining other in vivo methods.

In Vivo Microdialysis Method to Collect Large Extracellular Proteins from Brain Interstitial Fluid with High-molecular Weight Cut-off Probes

In vivo microdialysis has enabled collection of molecules present in brain interstitial fluid (ISF) from awake, freely-behaving animals. In order to analyze relatively large molecules in ISF, the current article specifically focuses on the microdialysis protocol using probes with high molecular weight cut off membranes.

In vivo microdialysis is a powerful technique to collect ISF from awake, freely-behaving animals based on a dialysis principle. While microdialysis is an ...

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is centered on culturing small pieces of embryonic or early postnatal rodent brains inside a 3-D hydrogel formed by the rat tail tendon-derived type I collagen with specific porosity. Tissue pieces are cultured inside the hydrogel and confronted to specific brain fragments or genetically-modified cell aggregates to produce and secrete molecules suitable for creating a gradient inside the porous matrix. The steps of this protocol are simple and reproducible but include critical steps to be considered carefully during its development. Moreover, the behavior of growing axons can be monitored and analyzed directly using a phase-contrast microscope or mono/multiphoton fluorescence microscope after fixation by immunocytochemical methods.

Here, we provide a method for analyzing the behavior of growing axons in 3D matrices, mimicking their natural development.

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is ...