Using Localization Microscopy to Quantify Calcium Channels at Presynaptic Boutons

Calcium channels at synaptic boutons are critical for synaptic function, but their number and distribution are poorly understood. This gap in knowledge is primarily due to the resolution limits of fluorescence microscopy. In the last decade, the diffraction limit of light was surpassed, and fluorescent molecules can now be localized with nanometer precision. Concurrently, new gene editing strategies allowed direct tagging of the endogenous calcium channel genes—expressed in the correct cells and at physiological levels. Further, the repurposing of self-labeling enzymes to attach fluorescent dyes to proteins improved photon yields enabling efficient localization of single molecules. Here, we describe tagging strategies, localization microscopy, and data analysis for calcium channel localization. In this case, we are imaging calcium channels fused with SNAP or HALO tags in live anesthetized C. elegans nematodes, but the analysis is relevant for any super-resolution preparations. We describe how to process images into localizations and protein clusters into confined nanodomains. Finally, we discuss strategies for estimating the number of calcium channels present at synaptic boutons. Key features • Super-resolution imaging of live anesthetized C. elegans. • Three-color super-resolution reconstruction of synapses. • Nanodomains and the distribution of proteins. • Quantification of the number of proteins at synapses from single-molecule localization data.


Background
Proteins' scale is in the order of nanometers; for example, GFP is approximately 4 nm across.However, due to the diffraction limit of light, collecting and focusing the light from GFP onto a camera sensor will, at best, produce an Airy disk with a diameter of 440 nm.To understand the distribution of proteins in cells, microscopy methods that can bypass the diffraction limit of light are required.Resolution is the spatial limit at which the certainty of resolving two nearby proteins fails; thus, high spatial resolution becomes particularly important for small subcellular structures like synapses [1].Synapses contain highly specialized domains, such as the active zone, but are just a few hundred nanometers in diameter.These highly organized domains allow rapid and reliable neurotransmission [2][3][4].To understand how these domains work, the underlying proteins and their nanoscale organization relative to each other must be determined.Super-resolution methods that achieve resolutions below 100 nm can be divided into stimulated emission microscopy (for example, STED) or localization microscopy (for example, PALM and STORM).Stimulated emission is a scanning method that, in essence, reduces the diameter of the laser beam.Localization microscopy illuminates the whole sample and collects a 2D image of fluorescence on a camera face but limits the number of molecules that contribute to any single frame of the movie so that they appear as single blinks of light.Next, a modeled 3D-point spread function is fit to the blinks of light to reconstruct an image in x, y, and z [5][6][7][8].The coordinate data produced from fitting a 3D-point spread function to the signal from each emitter has a precision determined by the Cramér-Rao lower bound (reviewed in von Diezmann et al. [9]).In contrast, the entire set of coordinate data achieves a resolution that can be determined by several methods.Fourier ring analysis is one common method to determine the resolution that reflects both localization precision and how well a set of localizations samples the underlying structure [10].By this metric, we typically obtain datasets with 40 nm resolution from live worms.Given that ion channel complexes are ~10 nm in diameter, these data fall short of resolving single channels.Beyond single proteins, we aim to determine the organization of two different proteins relative to one another.The spatial distributions of distinct proteins can be related in four ways: the proteins can be organized into a single coincident domain, overlapping domains, or adjacent domains, or be completely disjointed.Critically, the borders of protein domains, like the borders of a country, describe the essential information.While more difficult to analyze than an image, coordinate data can more completely describe these spatial relationships.Here, we demonstrate multicolor super-resolution imaging in live anesthetized C. elegans of directly tagged and labeled endogenous proteins.Further, we discuss quantifying proteins and analyzing images from 3D singlemolecule localization microscopy data.This protocol describes how to collect and analyze data from individual 3 Published: Aug 20, 2024 synapses, to count the number of ion channels, and determine the spatial relationships between nanodomains of different proteins.For counting single proteins, the use of endogenous labels vs. antibodies has two advantages.The first is saturation.The TMR dye saturates endogenous proteins tagged with HALOtag in one round of staining (Figure 1).Therefore, the number of proteins can be quantified without caveats from epitope availability or binding affinities of antibodies [11].The second is precision.Antibodies introduce an additional ~20 nm of uncertainty into localization due to their size.By contrast, the size of self-labeling enzyme tags is approximately 4 nm.One drawback of live cell imaging vs. antibodies on fixed samples is that oxygen-scavenging buffers cannot be used, leading to increased photobleaching (and thus lower photon counts or fewer localizations).Given these considerations, our protocol produces a reconstruction with sufficient localizations and resolution considering the typical size and separation of neuronal protein microdomains [12][13][14].
Figure 1.HALO-TMRstar saturates UNC-10::HALO after 60 min.L4 animals were incubated with 5 μM HALO-TMRstar (Green) for 60 min.After initial staining, animals were allowed to recover on a nematode growth media (NGM) plate with OP50 bacteria for 1 h.The animals were stained again in 5 μM HALO-SIR (Red) for 60 min and then allowed to recover for 4 h.Synaptic regions near the pharynx were imaged by localization microscopy.Scale bar = 1 μm.
To determine if two protein microdomains are associated, Pearson's or Mandel's correlation coefficients are commonly used to determine colocalization in fluorescence microscopy [15].These tests rely on diffraction-limited signals to create overlapping signals.However, for super-resolution methods, as spatial resolution approaches the size of a protein, the correlation coefficients in these tests will approach zero.For these reasons, it is preferable to analyze the data using nearest neighbor values.Importantly, nearest neighbor analysis distinguishes domains that are coincident, domains that do not overlap but are adjacent, domains at fixed distances, and domains with no relationship.In contrast to nearest neighbor measurements, the spatial relationship of localizations to a center axis or biological landmark has been successful at describing the underlying biology [1,16].In the following protocol, we describe how to count the number of proteins in a nanodomain, how to measure the size of the domain, and how to determine relationships between protein domains based on localization coordinates.For this study, we used a Bruker Vutara SRX352.In brief, this is a super-resolution microscope that uses wide-field illumination of the sample.Wide-field illumination activates fluorophores and collects light from an entire vertical column; thus, thin samples or samples positioned with regions of interest near the objective are crucial for reducing background emissions.The microscope uses a proprietary optical biplane that splits the focal plane by ~700 nm in Z-space onto the camera face.Biplane imaging permits improved axial resolution during reconstructive microscopy and provides 1.2 μm of optical thickness at a single z-position.Further, this microscope is capable of 3-color superresolution imaging with 488, 561, and 641 nm excitation lines.These laser lines are paired with emissions filters for green (BP497-538), orange (BP570-629), and red (BP647-739) fluorophore imaging.Although we describe the specific routine for a Vutara SRX352, in principle the steps listed here are applicable to any super-resolution microscope.
Here, Sections 1-4 deal with the details for mounting and imaging C. elegans nematodes, and Sections 4-8 concern imaging and analysis of localizations.

B. Label SNAP and HALO
1. Day 4, evening a. Inspect plates to select for broods with an abundance of L4 hermaphrodite animals.This is a synchronized plate.b.Wash animals off the three synchronized plates using ~1 mL of M9 buffer and a glass Pasteur pipette.
Place animals into a microcentrifuge tube.Worms will stick to the plastic of a micropipette, so use glass to move animals.c.On a benchtop centrifuge, gently spin (10 s each) to remove bacteria, removing the supernatant and washing with 1 mL of M9 buffer.

C. Imaging preparation
1. Day 5, morning a.Prepare a 4% agarose pad by heating an agarose aliquot in a 95 °C heat block and placing a small drop on a microscope slide.Then, flatten the pad with another microscope slide (Figure 2A).b.Trim the agarose pad to about 10 mm × 5 mm using a microscope slide.The agarose pad should fit entirely under the coverslip (Figure 2B).c.Add 2-4 μL of 25 mM NaN3 to the pad.d.Use the worm pick to distribute the NaN3 drop evenly across the agarose pad.If NaN3 flows over the edges of the agarose pad, reduce the amount used.e. Transfer 20 young adult worms to the agarose pad.Older animals may have been young adults during staining and will be highly fluorescent because they have not shed their cuticle.f.Wait for animals to paralyze and straighten.This should take a few minutes.If the droplet of NaN 3 is drying out, place the slide into a humidity chamber.Seal the pad with a coverslip.Then , seal the coverslip with nail polish and image (Figure 2B).

D. Superresolution imaging of C. elegans
1. Image worms by 3-color imaging a. Organize laser lines in descending order of wavelength, e.g., 1) 646 nm, 2) 549 nm, 3) 488 nm.This reduces the photobleaching of the sample.b.In widefield mode: Locate worms on the pad and coverslip.c.In super-resolution mode: i. Use only extremely low laser powers (< 1%).ii.Set the exposure time to 300 ms.
iii.Focus both imaging planes as much as possible on the dorsal nerve cord (Figure 2A).iv.Turn lasers off as soon as possible.We try to have the laser on for under 10 s while focusing on the synapses we plan to image.d.After the nerve cord is in focus, reduce the exposure time to 30 ms and proceed to record.Tune laser powers for optimal blinking vs. the lifespan of the fluorophores.An example of a properly tuned laser power is shown in Figure 3B.Notes: i.The best samples are worms that are oriented with their dorsal nerve cord tilted toward the objective.
ii. Be very cautious with the laser power while finding your sample.The worm is not in oxygen scavenging buffer; thus, fluorophores will bleach.To focus on the cord, use very low laser power and a 300 ms exposure time.
iii.Optimizing laser power while imaging: A laser power that is too low will not induce blinking of the fluorophores.A laser power that is too high will bleach the sample in a few frames.The user should find these limits and note them for their microscope.The localization counts over time displayed in Figure 3B is an example of an ideally tuned laser power.

Image worms-counting channels
a. Organize laser lines by starting with the protein you wish to quantify, followed by fiducial markers.Fiducial markers label a known structure, like the synaptic dense projection, and are used to spatially orient the experimental channel.b.In widefield mode: Locate worms on the pad and coverslip.c.In super-resolution mode: i. Use only extremely low laser powers (< 1%).ii.Set the exposure time to 300 ms.
iii.Focus both imaging planes as much as possible on the dorsal nerve cord.iv.Turn lasers off as soon as possible.d.After the cord is in focus, reduce the exposure time to 30 ms and proceed to record.Increase the laser powers for optimal blinking vs. lifespan of the fluorophores.This is typically between 4% and 12% with 200 mW lasers and the 40 μm biplane module.For counting emitters, the dyes should be completely bleached after imaging; this typically occurs within 4,000 frames.Notes: i.The best samples are worms that are oriented with their dorsal nerve cord tilted toward the objective.ii.Optimizing laser power while imaging: A power that is too low will not induce blinking of the fluorophores.A laser power that is too high will bleach the sample in a few frames.The user should find these limits and note them for their microscope.The localization counts over time displayed in Figure 3B is an example of an ideally tuned laser power.

E. Localizing
1. Proceed to the Localization tab.
2. Set the background threshold.This should be optimized to minimize background localizations (false positive) but not exclude real emitters (false negative).Adjust the background threshold to produce cutouts (colored boxes) around real signals but minimal cutouts in areas with no signal.An example of the effect

F. Visualizing
Proceed to the Visualization tab. 1. Discard localizations that are not accurate in the Advanced Particle Filters dialogue.For live worms, we set an arbitrary threshold of 50 nm radial precision and display localizations on the visualization plot as a 50 nm particle size (Figure 4A). 2. Denoise the image using Method: Mean Distance and denoise by Percent.Lower the slider until most of the signal outside the cord is removed but the signal inside the cord appears unperturbed (Figure 4B). 3. Save this image to the View Manager.4. For analysis of proteins at individual synaptic boutons, select a region of interest 700 nm radius from the center of an in-focus dense projection.Save this view in View Manager.Export the data and save it as a particle .csvfile.Out-of-focus synapses and synapses with chromatic aberrations in the XY plane are discarded.Some chromatic aberrations will be present in the Z plane.Aim to analyze five synapses per animal from five different animals, with a total of 25 synapses analyzed.We recommend saving particle files ordered by "animal#_synapse#", e.g., "3_1.csv."

Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):

ProcedureA. 2 .
Synchronize worms by staging embryos1.Day 1 a.At 9:00 am, use a worm pick to move 20 young adult animals onto a new NGM plate seeded with OP50; do this three times to create three plates total.This procedure takes approximately 10 min.b.At 5:00 pm, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three additional plates.Days 2-3 a.At 9:00 am, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three additional plates.b.At 5:00 pm, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three new plates.c.Move the original adult animals to fresh plates in the morning and evening for one more day.
d.After three washes, remove the supernatant and resuspend the animals in approximately 195 μL of M9 buffer.e. Prepare JF dyes by resuspending at 1 mM in DMSO (e.g., 5 nmol of dye in 5 μL of DMSO).Move dye + DMSO suspension into the next dried dye aliquot to create a combination of dyes in suspension.This will create 5 μL of 1 mM JF646-HALOtag Ligand + JF549cp-SNAPtag Ligand dye.f.Add dye + DMSO solution to worms in M9 buffer to create a 25 μM final concentration of dye.Mix gently by vortexing.Note: DMSO above 5% is toxic to worms.g.Put the tube with the lid in a freezer box on an orbital shaker and stain for 120 min at RT. Gently flick and spin down worms every 15 min to prevent worms from forming clumps on the bottom of the tube or becoming stuck on the sides of the tube.It is essential that the dye and worms incubate in a dark place like a lidded freezer box.h.After 2 h, spin down the worm solution, aspirate off the supernatant, and wash four times with 1 mL of M9 buffer.i.With a Pasteur pipette, transfer worms to at least three OP50-seeded NGM plates.Let the M9 droplet dry completely before inverting the plate.This should be performed in the dark (we use a drawer).After the droplet has dried, cap the plates and invert.Let the worms recover in a lidded freezer box for 10-12 h.It is important to not expose the stained worms to light.Ideally, the L4 to adult molt will occur during recovery to minimize background staining of the cuticle and gut.Note: L4 hermaphrodite animals can be identified by the absence of a vulva, which appears as a white crescent at the center of the ventral side of the animal under a typical dissection microscope.Starting with synchronized plates is perhaps the most crucial step of the protocol because the dye sticks to the cuticle of the animal.If adults are exposed to dye, they will have non-specifically labeled cuticles forever, which interferes with imaging of the nervous system.L4s are the final larval stage of the animal and must molt their cuticle before adulthood.Thus, if L4s are stained and then imaged as young adults, a molt must have occurred during recovery.

7 Published
Cite as: Mueller, B.D. et al. (2024).Using Localization Microscopy to Quantify Calcium Channels at Presynaptic Boutons.Bio-protocol 14(16): e5049.DOI: 10.21769/BioProtoc.5049.not image animals that have been in NaN3 for more than 60 min.The paralytic sodium azide causes neuron blebbing and disruption of cell morphology after approximately 1 h.ii.Avoid air bubbles in the agarose pad or trim them.

Figure 2 .
Figure 2. Creating a worm imaging pad and slide.(A) The agarose pad acts as a support for the worm sample so it is not crushed by the microscope slide, also immobilizing the animal.(B) Once sealed with nail polish, the worms should last for hours.However, the sodium azide paralytic causes cell blebbing and disruption of cell morphology after about an hour.(C) Left: Under the brightfield illumination of the Vutara SRX 352, the worms will appear as pictured.Right: The roll can be in part determined by the position of the ventral nerve cord exiting the neuropil if there is a fluorescent marker in the nervous system or position of the vulva.Scale bar = 5 μm.

9 Published:Figure 3 .
Figure 3. Biplane, blinking, and background thresholding.(A) The Vutara SRX simultaneously images in two focal planes ("biplane"), which splits the light path into two focal planes separated by about 700

Figure 4 .
Figure 4. Data filtering.(A) First, reconstructions are filtered to exclude imprecise localizations.(B) Next, localizations are trimmed based on nearest neighbors using the denoising tool.Finally, a region of interest is drawn around a synapse to be analyzed to exclude localizations that cannot be within the synapse based on the size of the bouton.Numbering and green arrows show the click order for the user.Magenta = CaV2, Yellow = CaV1.

: Aug 20, 2024 3. 4% agarose in M9
Heat in the microwave until dissolved.Store at 4 °C in aliquots to avoid evaporation from heating cycles.Agarose aliquots have a shelf-life of one year.Dispose if evaporation or fungal or bacterial contamination occurs.Benchtop centrifuge (Thermo Fisher, model: mySPIN6, catalog number: 75004061) 3. Heat block set to 95 °C (e.g., Eppendorf, model: ThermoMixer F1.5, catalog number: 5384000020) Click on Advanced Statistics Dialogue and go to the Cluster Analysis tab. 2. Set the parameters for calling clusters based on the size of the protein and resolution of the image.See note.3. Click Compute. 1.