Improved methods for single-molecule fluorescence in situ hybridization and immunofluorescence in Caenorhabditis elegans embryos

Abstract

Visualization of gene products in Caenorhabditis elegans has provided insights into the molecular and biological functions of many novel genes in their native contexts. Single-molecule Fluorescence In Situ Hybridization (smFISH) and Immunofluorescence (IF) enable the visualization of the abundance and localization of mRNAs and proteins, respectively, allowing researchers to ultimately elucidate the localization, dynamics, and functions of the corresponding genes. Whereas both smFISH and immunofluorescence have been foundational techniques in molecular biology, each protocol poses challenges for use in the C. elegans embryo. smFISH protocols suffer from high initial costs and can photobleach rapidly, and immunofluorescence requires technically challenging permeabilization steps and slide preparation. Most importantly, published smFISH and IF protocols have predominantly been mutually exclusive, preventing the exploration of relationships between an mRNA and a relevant protein in the same sample. Here, we describe protocols to perform immunofluorescence and smFISH in C. elegans embryos either in sequence or simultaneously. We also outline the steps to perform smFISH or immunofluorescence alone, including several improvements and optimizations to existing approaches. These protocols feature improved fixation and permeabilization steps to preserve cellular morphology while maintaining probe and antibody accessibility in the embryo, a streamlined, in-tube approach for antibody staining that negates freeze-cracking, a validated method to perform the cost-reducing smiFISH (single molecule inexpensive FISH) adaptation, slide preparation using empirically determined optimal antifade products, and straightforward quantification and data analysis methods. Finally, we discuss tricks and tips to help the reader optimize and troubleshoot individual steps in each protocol. Together, these protocols simplify existing workflows for single-molecule RNA and protein detection. Moreover, simultaneous, high-resolution imaging of proteins and RNAs of interest will permit analysis, quantification, and comparison of protein and RNA distributions, furthering our understanding of the relationship between RNAs and their protein products or cellular markers in early development.

INTRODUCTION:

The study of spatial and temporal expression patterns can provide fundamental insights into gene function and importance. By querying the abundance and spatial patterning of mRNAs and their protein products in whole animals, it is possible to gain an understanding of their transcription, translation, mRNA stability, protein dynamics, developmental regulation, and their functional roles (Levesque & Raj, 2013; Parker et al., 2020; Perez-Burgos et al., 2003; Toki et al., 2017; Trcek et al., 2017). Visualizing RNAs and proteins in the same intact animal requires methods that are sensitive, minimally disruptive, and, most importantly, compatible with one another. Typically, researchers label target RNAs and proteins in situ, using single-molecule Fluorescence In Situ Hybridization (smFISH) and immunofluorescence (IF), respectively. In smFISH, hybridization of fluorescent antisense probes labels RNAs, and in IF, fluorescent antibodies stain proteins (Femino et al., 1998; Orjalo et al., 2011; Raj et al., 2008). Thus, researchers can study the relationships between RNA and proteins by performing smFISH and IF protocols on the same sample.

Standard smFISH and IF protocols, however, do have challenges. For instance, low signal plagues standard smFISH protocols, due to the poor photostability of some fluorophores. High background exacerbates this low signal as a result of the relatively thick z-dimension of C. elegans embryos (Ji & Oudenaarden, 2012). These issues ultimately lead to low signal-to-noise ratios (SNR) and difficulty in detecting RNA spots. Further, the probes used in the standard smFISH protocol are costly, preventing many research groups from adopting this technique. Meanwhile, standard IF protocols pose problems in C. elegans embryos (Tsanov et al., 2016) due to the embryo’s strong eggshell and robust permeability barrier (J. Duerr, 2006; Olson et al., 2012). Ultimately, this has resulted in the use of harsh fixatives (aldehydes, picric acid), reducing reagents (B-mercaptoethanol, DTT), and enzymatic treatments (collagenase), which are capable of damaging nucleic acids (Bonin et al., 2005; J. S. Duerr, 2013; Hoetelmans et al., 2001; Masuda et al., 1999; Tyrrell et al., 1995; Williams et al., 1999), and require technically-demanding slide preparations for freeze-crack permeabilization (J. Duerr, 2006; J. S. Duerr, 2013). Perhaps most importantly, most smFISH and IF protocols are mutually exclusive in the C. elegans embryo due to incompatibilities in the reagents and protocols used.

By optimizing, simplifying, and combining smFISH and IF protocols, we have remedied several of the issues in existing protocols. In Basic Protocol 1, we present a one-tube protocol to perform IF and smFISH sequentially on the same sample. In this protocol, embryos are first harvested, fixed, and permeabilized using mild conditions, thus mitigating issues of nucleic acid damage and challenging slide preparations. IF is then performed on fixed embryos using reagents that protect the integrity of RNA in the sample, allowing smFISH to be performed subsequently. Finally, slides are prepared using antifade solutions that protect the smFISH signal from photobleaching and reduce the background signal to raise the SNR of RNA spots. In Alternate Protocol 1, an abbreviated protocol for performing IF and smFISH is described, where embryos are harvested, fixed, and a standard smFISH protocol is performed with the inclusion of nanobodies to stain protein. For those interested in performing the techniques individually, we provide two protocols. Basic Protocol 2 describes a simplified immunofluorescence protocol, and Basic Protocol 3 describes the steps to perform smFISH, including steps to perform the low-cost adaptation, single-molecule inexpensive Fluorescence In Situ Hybridization (smiFISH). An overview of the entire workflow is shown in Figure 1.

Figure 1. Schematic illustration of the IF, FISH, and IF/FISH protocols described in this article.

An overview illustrating the workflow of the sequential IF/FISH (Basic Protocol 1), simultaneous IF/FISH (Alternate Protocol 1), simplified IF (Basic Protocol 2), and smFISH or smiFISH (Basic Protocol 3), from sample preparation to slide preparation.

It is worth noting that while we will discuss some data analysis tools and best practices, we will not describe here how to perform data analysis in detail.

BASIC PROTOCOL 1: Sequential Immunofluorescence and single-molecule Fluorescence In Situ Hybridization

This protocol describes methods for isolating C. elegans embryos and fixing them in a manner compatible with both immunofluorescence and RNA FISH. Steps for performing immunofluorescence subsequently followed by smFISH are then outlined. Finally, slide preparation is described. This approach can be used for simultaneous visualization of RNA transcripts and a protein of interest, provided the FISH probes and fluorescent antibody are selected in distinct channels.

CAUTION: To avoid RNA degradation, use RNA-appropriate sterile technique and RNAse-free reagents where possible.

CAUTION: Many reagents in this protocol are hazardous. Follow all appropriate guidelines and regulations for the use and disposal of each reagent.

Materials:

C. elegans strain/genotype of interest (Caenorhabditis Genetics Center, CGC)

Nematode Growth Media Plates (see recipe in Reagents and Solutions)

OP50 E. coli (Caenorhabditis Genetics Center, cat. no. OP50)

100% Reagent Grade Acetone (Fisher, cat. no. A18–500), cooled to −20 °C

100% Reagent Grade Methanol (Fisher, cat. no. A412–500), cooled to −20 °C

Bleaching Solution (see recipe in Reagents and Solutions)

M9 buffer (see recipe in Reagents and Solutions)

10X PBST (see recipe in Reagents and Solutions)

Bovine Serum Albumin (Sigma, cat. no. A9418–5G)

Primary Antibody or Fluorescently Labeled Nanobody/ScFv e.g. K76, anti-PGL-1 (DHSB, Cat# K76, RRID:AB_531836)

Fluorescent Secondary Antibody, e.g. Alex Fluor 488 Goat Anti-Mouse IgG (H+L) secondary antibody (Jackson ImmunoResearch, cat. No. 115–545-03, RRID: AB_2338840)

DAPI, 4’,6-Diamidino-2-Phenylindole, Dihydrochloride (Invitrogen, cat. no. D1306)

RNasin^® Ribonuclease Inhibitor (Promega, cat. no. N2111)

2X SSC (see recipe in Reagents and Solutions)

Wash Buffer A (see recipe in Reagents and Solutions)

Wash Buffer B (see recipe in Reagents and Solutions)

Hybridization Buffer (see recipe in Reagents and Solutions)

Mounting Medium (see recipe in Reagents and Solutions)

smFISH probes, 25 uM stocks in 10 mM Tris pH 8.0, e.g. nos-2 probe set (Biosearch Technologies, Custom Probes made using the Stellaris Probe Designer)

RNAse-free Water (Invitrogen, cat. no. AM9922)

VECTASHIELD Mounting Medium (Vector Laboratories, cat. no. H-1000–10)

Liquid Nitrogen

150 mm Petri Dishes (Genesee Scientific, cat. no. 32–106, or equivalent. Other sizes are acceptable.)

Fume Hood (Hamilton Safeaire II, cat. no. 54L2769, or equivalent)

Stereomicroscope (Nikon, cat. no. SMZ800N or equivalent)

10 mL Serological Pipets (Genesee Scientific, cat. no. 12–104 or equivalent)

25 mL Serological Pipets (Genesee Scientific, cat. no. 12–106 or equivalent)

Pipet Controller (Drummond, cat. no. 4–000-101 or equivalent)

2000 mL Erlenmeyer Flask (Sigma, cat. no. CLS51002L or equivalent)

1000 mL Glass Bottles (VWR, cat. no. 10754–820 or equivalent)

250 mL Glass Bottles (VWR, cat. no. 89000–236 or equivalent)

15 mL Centrifuge Tubes (Olympus Plastics, cat. no. 28–103 or equivalent)

1.5 mL Microcentrifuge Tubes (Olympus Plastics, cat. no. 14–125 or equivalent)

1.5 mL Amber Microcentrifuge Tubes (Olympus Plastics, cat. no. 22–282AM or equivalent)

Nutating Mixer (Labnet, cat. no. S0500 or equivalent)

Centrifuge (Eppendorf, cat. no. 5810 or equivalent)

Microcentrifuge (Eppendorf, cat. no. 5424 or equivalent)

Mini Centrifuge (Genesee, cat. no. 27–523 or equivalent)

Vortex Mixer (VWR, cat. no. 12620–838 or equivalent)

Liquid Nitrogen Container (Thermo Scientific, cat. no. 2122 or equivalent)

Thermal Cycler (Bio-Rad, cat. no. T100 or equivalent)

ThermoMixer (Eppendorf, cat. no. 05–412-503)

8 mm 1.5 thickness round cover glass (Electron Microscopy Sciences, cat. no. 72296–08)

Glass microscope slides (VWR, cat. no. 48312–401)

1.5 thickness, 22×22 mm cover glass (VWR, cat. no. 48366–227)

Grace Bio-Lab Press-To-Seal silicon isolator (Sigma, cat. no. GBL664504–25ea)

Widefield Fluorescent Microscope (DeltaVision Elite or equivalent)

−20 °C Freezer

4 °C Refrigerator

15–25 °C incubator

10, 200, and 1000 uL Micropipettes and tips

Protocol steps

Embryo Preparation and Fixation

• 1Grow worms to gravidity on OP50-seeded NGM plates. Synchronize by bleaching if necessary.

  Typically, one or two 150 mm NGM plates seeded with ∼2 mL of OP50, chunked with the strain of interest and grown until worms are gravid, is sufficient. Other bacterial stocks, such as inducible RNAi vector-containing E. coli, can be used if desired.

• 2Harvest gravid worms by washing them off plates using M9 and collecting them in a 15 mL conical tube in ∼15 mL total volume.

  Aggressive pipetting will increase yield by releasing more worms from the plates. Be sure not to pierce the plate’s surface, as agar carried into the sample will persist.

• 3Spin conical at 2000 × g for 1 min at room temperature to pellet gravid worms. Alternatively, allow gravid worms to settle over time.
• 4Remove supernatant using a pipette or aspirator, being careful not to disturb the worm pellet.
• 5Resuspend the worm pellet in 15 mL of M9.
• 6Repeat Step 3.
• 7Repeat steps 4 – 6 until the supernatant is clear, discarding the supernatant after the final wash.
• 8Add ∼15 mL of bleaching solution to the worms and nutate or hand-shake for 6–8 min until embryos are released from the mothers.

  Check on the condition of the worms periodically throughout bleaching. The gravid adults should break into about two pieces before continuing. If worms are bleached for too long, some early-stage embryos may be damaged. For tips on harvesting embryos, see (Porta-de-la-Riva et al., 2012).

• 9Centrifuge conical at 2000 × g for 1 min at room temperature to pellet. Immediately remove supernatant and quench bleaching with 15 mL of M9.

  At this point, embryos typically stick to the tube, and the supernatant can be carefully decanted to decrease the time before quenching.

• 10After adding M9, vortex the pellet to release remaining worm fragments. Centrifuge at 2000 × g for 1 min at room temperature.
• 11Discard M9 and wash with 15 mL of M9 two more times (for a total of 3 washes), vortexing the pellet after the addition of M9 each time.

  The aroma of bleach should be completely gone by the end of washing.

• 12Transfer the remaining embryos to a 1.5 mL microcentrifuge tube and pellet in a tabletop microcentrifuge for 30 sec at 2000 × g at room temperature. Turn tube 180° and repeat until a pellet has formed. Typically, between 2–4 spins is sufficient. Remove any remaining M9.

  C. elegans embryos will stick to the side of microcentrifuge tubes during centrifugation, leading to a smear of embryos and poor pelleting if tubes are not rotated and spun repeatedly.

• 13Add 1 mL of pure methanol cooled to −20 °C, vortex to break up the pellet, and immediately submerge in liquid nitrogen for 1 min to crack the eggshell and promote permeabilization.
• 14Remove the tube from the liquid nitrogen and immediately begin pelleting at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin.

  The sample will still be partially frozen for the first spins, but it is best to get the sample pelleting early to prevent over-fixation.

• 15Repeat Step 14 until the embryos are pelleted, or until the embryos have been in methanol for a total of 5 min.
• 16Once the sample has been in methanol for a total of 5 min, including the time spent pelleting, remove the methanol and replace it with 1 mL of pure acetone cooled to −20 °C. Store the sample at – 20 °C for ∼3 min.
• 17Pellet embryos by centrifugation as in step 14 until a pellet forms or the embryos have been in acetone for a total of 5 min.
• 18After embryos have fixed in acetone for a total of 5 min, remove the acetone and immediately continue to immunofluorescence.

Immunofluorescence

• 19Add 1 mL of 1X PBST to the sample from Step 18 and nutate for 5 min to wash.
• 20Pellet embryos by centrifuging at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until a pellet forms.
• 21Pipet out or aspirate as much of the supernatant PBST as possible without disrupting the pellet.
• 22Repeat steps 19–21 two more times (3 washes total).
• 23Add 250 uL of 1X PBST containing 1% w/v BSA. Incubate for 30 min at 37 °C with nutation, to block.

IMPORTANT: If RNA FISH will be performed subsequently, it is essential to add 1 unit/uL RNasin^® (Promega) during steps where BSA is included, to prevent RNA degradation. RNAse-free BSA can be used if issues with RNA degradation occur with sequential IF/smFISH protocols; however, it is much more expensive.

• 24Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 25Pipet or aspirate as much of the supernatant as possible without disrupting the pellet.
• 26Add a maximum of 100 uL of the primary antibody diluted in 1X PBST with 1% w/v BSA (and 1 unit/uL RNasin^® if FISH will be performed subsequently). Nutate at room temperature for at least 1 hr, or overnight at 4 °C.

  Example: For the data shown in Figure 2, we used 40 ul of a 1:20 dilution of K76 antibody and 100 ul of a 1:1000 dilution of 2A4 antibody.

  Overnight incubations provide better IF signals but can increase RNA degradation. Optimal antibody concentrations must be determined for each antibody. The volume of primary antibody solution added should be minimized to reduce the consumption of antibodies, where possible.

• 27Add 1 mL of 1X PBST directly to the sample and nutate for 5 min to wash out free antibody.
• 28Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until a pellet forms.
• 29Pipet out or aspirate as much of the supernatant PBST as possible without disrupting the pellet.
• 30Repeat steps 27–29 two more times (3 washes total).
• 31Add a maximum of 250 uL of fluorescently labeled secondary antibody diluted in 1X PBST and incubate for at least 1 hr in the dark at room temperature with nutation.

  Example: For the images in Figure 2, we used 125 ul of a 1:250 dilution of Alex Fluor 488 Goat Anti-Mouse secondary antibody.

  Optimal antibody concentrations must be determined for each antibody. The volume of secondary antibody solution added should be minimized to reduce the consumption of antibodies, where possible.

• 32Add 1 mL of 1X PBST and nutate for 5 min to wash out excess antibody.
• 33Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 34Pipet out or aspirate as much of the supernatant PBST as possible without disrupting the pellet.
• 35Repeat steps 32–34.
• 36Add 1 mL of 2X SSC and nutate for 5 min.

  2X SSC facilitates smFISH probe hybridization. Washing in 2X SSC is used to remove PBST and equilibrate embryos in an smFISH-compatible solution.

• 37Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 38Pipet out or aspirate as much of the supernatant 2X SSC as possible without disrupting the pellet.
• 39Repeat steps 36–38.
• 40Continue to smFISH.

Figure 2. Sequential IF/FISH.

Immunofluorescence followed by smFISH was performed on N2 embryos. IF was performed using K76 (A and B) or 2A4 (C) primary antibodies to identify PGL-1-containing P granules and ELT-2 protein (magenta), respectively. smFISH was used to simultaneously detect the P granule constituent RNAs nos-2 (A) and cpg-2 (B), or elt-2 mRNA (C), all in magenta. Embryos were counterstained with DAPI (blue). Three biological replicates were performed for each experiment. Scale bars represent 10 μm.

smFISH

• 41Prepare fresh buffers by adding 10% formamide to stocks of Wash Buffer A and Hybridization Buffer. For each sample, prepare separate tubes with 3 ml of Wash Buffer A and 110 ul of Hybridization Buffer, and add formamide to each to a final concentration of 10%,.

  Wash Buffer A and Hybridization Buffer should always have formamide added immediately preceding the experiment. Formamide can decompose over time, particularly at higher temperatures, leading to less stringent probe binding. It can also acidify when exposed to air resulting in fluorophore quenching. Formamide stocks should be stored frozen, and their pH monitored periodically (pH 7–8 is ideal)

• 42Prepare a 1:20 dilution of the 25 uM smFISH probes in RNAse-free water (i.e. 1.25 uM final concentration). Add 2 uL of the 1.25 uM dilution to 110 uL of hybridization buffer (with formamide). If performing experiments using multiple probe sets with different fluorophores, add 2 uL of each diluted probe set. Mix well. Hybridization buffer is viscous.

  Example: To obtain the data in Figure 2, we used 2 uL of 1.25 uM nos-2 (Quasar 670), cpg-2 (Cal Fluor 610), and elt-2 (Cal Fluor 610) probe dilutions.

  Although 2 uL of 1.25 uM probe solution has worked well for most of the probe sets we have used, it is helpful to perform a titration over ∼1 order of magnitude of concentrations (~0.25 uM – 2.5 uM) to identify optimal probe concentrations on an individual-probe-set basis.

• 43Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 44Pipet out or aspirate as much supernatant as possible without disturbing the pellet.
• 45Add 1 mL Wash Buffer A (with formamide) to prehybridize sample and incubate at room temperature for ∼5 min.

  Nutation is optional during this wash step.

• 46Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 47Pipet out or aspirate as much supernatant as possible without disturbing the pellet.
• 48Add 100 uL of the hybridization buffer with probes (from Step 42) to the pelleted embryos and hybridize at 37 °C in the dark for a minimum of 8 hours, shaking at 450 rpm to ensure even probe penetration.

  The standard Stellaris RNA FISH protocol suggests hybridization should occur for anywhere between 4–16 hours. We have found that hybridizations shorter than 8 hours lead to variable smFISH signals. Additionally, we have had successful experiments using hybridizations as long as 48 hours. It is worth noting that longer hybridizations can increase the risk of RNA degradation if the sample was contaminated with RNase during the immunofluorescence steps. Store prepared Wash Buffer A at room temperature or 37 °C during this incubation. Warm buffer will increase the stringency of probe binding and decrease background and non-specific binding. If available, use a thermomixer to shake the hybridization solution, although standard incubators are also acceptable.

• 49Add 1 mL of warm Wash Buffer A (with formamide) directly to the embryos in the hybridization solution.
• 50Incubate at 37 °C in the dark for 30 min, shaking at 450 rpm in a thermomixer or incubator.
• 51Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 52Pipet out or aspirate as much supernatant as possible without disturbing the pellet.
• 53Prepare 1 mL of Wash Buffer A (with formamide) containing 1 ng/uL DAPI for each sample to be imaged. Add 1 mL of the mix to the sample.

  If 3 ml of Wash Buffer A were prepared, as described in step 41, this should be the remaining volume of Wash Buffer A. No Additional buffer should need to be prepared.

• 54Incubate at 37 °C in the dark for 30 min, shaking at 450 rpm in a thermomixer or incubator.
• 55Centrifuge embryos at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.
• 56Pipet out or aspirate as much supernatant as possible without disturbing the pellet.
• 57Add 1 mL of Wash Buffer B and incubate for ∼5 min.
• 58Repeat step 55 and 56.
• 59Resuspend in 50 uL of mounting medium and incubate at 4 °C for 30 min to ensure antifade penetrance.

  Less mounting medium can be added if the embryo yield is low. The ideal volume of mounting medium to add is based on slide preparation. When a second cover glass is added to the sample in Step 63, 2–6 uL of embryos in mounting medium mixed 1:1 with VECTASHIELD should be dense enough that they will be easy to find on a microscope but sparse enough that they do not clump together.

• 60Move to slide preparation.

Slide Preparation

• 61Working at a dissecting microscope, drop 2–6 uL of embryos suspended in mounting medium (from Step 59) onto a single 8 mm 1.5 thickness round cover glass resting on a glass slide.

  The volume of embryos in mounting medium to add can be tuned to optimize slide preparation. Add enough so that the embryos will be abundant but not clumped together after adding an equal volume of VECTASHIELD in Step 62. Do not add more than 6 ul, as most of the sample will be squeezed out when the second cover glass is added

  Always wear gloves when handling slides and cover glass to prevent smudging and contamination.

• 62Add an equal volume of VECTASHIELD antifade to the embryos from Step 61 in mounting medium on the cover glass and pipet up and down to mix thoroughly. Try to keep the final volume on the cover glass to ∼4–6 uL by removing some of the mixture.

  This is a good time to break up any large clumps of embryos by pipetting.

• 63Place a 1.5 thickness 22 mm × 22 mm square cover glass on top, trying to avoid bubbles.

  Do not let the cover glass touch the slide. The sample solution will pour over the edge of the round cover glass and seal it to the slide beneath through surface tension. Having the round cover glass close to the edge of the slide can provide some extra working height. Additionally, gently lowering the square cover glass from front to back over the round cover glass until surface tension pulls the round cover glass up will help prevent spillover.

• 64Flip the cover glass sandwich so the square cover glass is on the bottom. Remove as much liquid as possible from between the two cover glasses using a torn Kimwipe placed against the round one.

  The aim is to flatten the embryos as much as possible without damaging them. Samples can be firmly pressed on with a pipette tip as long as the cover glass doesn’t slide from side to side. The ideal depth of an embryo on the slide is ∼12–20 um. The signal-to-noise ratio will decrease and photobleaching will increase, with increasing thickness due to out-of-focus light and more image acquisitions, respectively.

• 65Affix the cover glass sandwich to a microscope slide using a Grace Bio-Lab Press-To-Seal silicon isolator such that the embryos will be imaged through the square cover glass.
• 66Head off to the microscope.

Acquisition settings will vary depending on the molecules being imaged and the microscope set up. Using our GE DeltaVision Elite microscope with an Olympus PLAN APO 60X, 1.42 NA objective, an Insight SSI Solid State Light Engine, and the standard DeltaVision DAPI, FITC, mCherry, Cy5 polychroic filter set, our image acquisition settings are as follows for the sample data shown in Figure 2A and 2B: Cy5 (imaging nos-2 RNA labeled with Quasar 670), % transmittance = 100%, exposure = 0.8 sec; mCherry (imaging cpg-2 RNA labeled with Cal Fluor 610), % transmittance = 100%, exposure = 1.0 sec; FITC (imaging PGL-1 protein stained with K76 primary antibody and Alexa Fluor 488 secondary antibody), % transmittance = 10%, exposure = 0.025 sec; DAPI (imaging DNA), % transmittance = 2%, exposure = 0.025 sec. We always image through the entire Z-stack at the longest wavelength before moving to the next longest wavelength, to prevent photobleaching of the more labile fluorophores at the red end of the spectrum. We use 0.2 um z-spacing between images. Typically, imaging 10 embryos at a particular developmental stage over 3 biological replicates is sufficient to detect statistically significant changes in RNA and protein abundances or distributions. It is often worthwhile, however, to perform a power analysis using the measurements from a pilot experiment to determine requisite sample sizes empirically before performing true experimental replicates. It is essential to tightly bin embryos of different cell stages or use equal numbers of embryos from each stage for all conditions where statistical analysis will be performed. The RNA and protein content of embryos changes rapidly and can lead to skewed statistics if stages are not carefully monitored. For example, the abundance of the cpg-2 RNA changes from an average of ~12,500 molecules at the 2-cell stage to ~5,000 molecules at the 4-cell stage (Parker et al., 2020).

ALTERNATE PROTOCOL 1: Abbreviated Protocol for Simultaneous Immunofluorescence and single-molecule Fluorescence In Situ Hybridization

In some instances, an abbreviated protocol allows the user to perform IF and smFISH simultaneously, negating the two-step approach described in Basic Protocol 1. This is the case, for instance, when using the anti-GFP nanobody (Chromotek, cat. no. gt-250). Though not all antibodies work with this streamlined approach, we have had success with high-affinity antibody derivatives such as nanobodies, single-chain variable fragments, and fragmented antibodies (Doshi et al., 2014). Alternate Protocol 1 has multiple advantages over Basic Protocol 1, namely, it is faster, requires fewer reagents, and avoids some reagents utilized in IF that contain RNases. In Alternate Protocol 1, IF and smFISH are performed simultaneously using an adapted smFISH protocol. Embryos are first harvested and fixed. An adapted smFISH protocol is then performed, including high-affinity antibody derivatives (e.g., nanobodies, single-chain variable fragments, or fragmented antibodies (Doshi et al., 2014)) in the RNA FISH buffers. This adapted smFISH protocol circumvents the need to perform a separate immunofluorescence protocol and reduces the time from sample preparation to imaging from 2–3 days to roughly 16 hours. Samples are then prepared on slides and are ready for imaging.

In our hands, small, high-affinity antibody derivatives, such as nanobodies, have improved the success of this simplified protocol. Moreover, we have only had success with primary staining using this protocol. Immunofluorescence using secondary antibody amplification during wash steps has not succeeded in our hands; however, a recent study suggests that some primary and secondary antibody combinations are likely compatible with this protocol (Tocchini et al., 2021). It is essential to verify that antibody staining is effective, by comparing protein localization to known distributions or using Basic Protocol 2. If a desired antibody or antibody pair does not work with Alternate Protocol 1, it is more likely to succeed using Basic Protocol 1. Regardless, it is worth trying Alternate Protocol 1 when testing new antibodies due to the protocol’s comparative simplicity.

Materials:

C. elegans strain/genotype of interest (Caenorhabditis Genetics Center, CGC)

Nematode Growth Media Plates (see recipe in Reagents and Solutions)

OP50 E. coli (Caenorhabditis Genetics Center, cat. no. OP50)

100% Reagent Grade Acetone (Fisher, cat. no. A18–500), cooled to −20 °C

100% Reagent Grade Methanol (Fisher, cat. no. A412–500), cooled to −20 °C

Bleaching Solution (see recipe in Reagents and Solutions)

M9 buffer (see recipe in Reagents and Solutions)

Fluorescently Labeled Nanobody/ScFv e.g. anti-GFP nanobody (Chromotek, cat. no. GT-250)

DAPI, 4’,6-Diamidino-2-Phenylindole, Dihydrochloride (Invitrogen, cat. no. D1306)

Wash Buffer A (see recipe in Reagents and Solutions)

Wash Buffer B (see recipe in Reagents and Solutions)

Hybridization Buffer (see recipe in Reagents and Solutions)

Mounting Medium (see recipe in Reagents and Solutions)

smFISH probes, 25 uM stocks in 10 mM Tris pH 8.0, e.g. nos-2 probe set (Biosearch Technologies, Custom Probes made using the Stellaris Probe Designer)

RNAse-free Water (Invitrogen, cat. no. AM9922)

VECTASHIELD Mounting Medium (Vector Laboratories, cat. no. H-1000–10)

Liquid Nitrogen

Equipment needs are identical to Basic Protocol 1.

Protocol steps:

• 1Perform embryo prep and fixation as in Basic Protocol 1, Step 1–18.

Do not perform any immunofluorescence steps. Move directly from embryo preparation and fixation to the smFISH protocol.

• 2
  Perform smFISH as in Basic Protocol 1, Steps 41–60, with the following exceptions:

  1. At Step 42, add the appropriate concentration of primary antibody or antibody derivative to the hybridization buffer.
     Example: To generate the data in Figure 3, we used a 2.37 ug/mL final concentration of anti-GFP nanobody (conjugated to Janelia Fluor 549) and 2 uL of a 1.25 uM dilution of nos-2 (Quasar 670) smFISH probes.
  2. In Step 50, increase the length of the second Wash A incubation from 30 min to 1 hour.
     If attempting a primary- and secondary-based antibody pair for this protocol, add the appropriate concentration of secondary antibody during the second Wash A incubation, at Step 49.
  3. Perform imaging and analysis as in Basic Protocol 1, Steps 61–66.

Figure 3. Simultaneous IF/FISH.

smFISH was performed on N2 embryos with the addition of anti-GFP nanobody to hybridization buffer. nos-2 mRNA (magenta) was probed using smFISH probes conjugated to Quasar 670. PATR-1::GFP (green) signal was visualized using 2.37 ug/mL Janelia Fluor 549 (Tocris 6147) conjugated anti-GFP nanobody (Chromotek, gt-250) (top). A no nanobody control is also shown (bottom). DNA was counterstained with DAPI (blue). Three biological replicates were performed for each experiment. Scale bars represent 10 μm.

See sample data in Figure 3.

BASIC PROTOCOL 2: Simplified Immunofluorescence in C. elegans embryos

This protocol presents a simplified protocol for performing IF alone in C. elegans embryos. It can be desirable to image protein distributions without visualizing RNA in the same sample, to simplify the procedures. Here, embryos will be harvested before being concurrently fixed and permeabilized using a combination of methanol and acetone. IF is then performed to stain a protein of interest. Finally, due to the optimized fixation steps, a simple slide preparation protocol can be used in place of the traditional freeze-cracking method. Ultimately, the user will then visualize the protein of interest on a fluorescent microscope.

Materials:

C. elegans strain/genotype of interest (Caenorhabditis Genetics Center, CGC)

Nematode Growth Media Plates (see recipe in Reagents and Solutions)

OP50 E. coli (Caenorhabditis Genetics Center, cat. no. OP50)

100% Reagent Grade Acetone (Fisher, cat. no. A18–500)

100% reagent grade methanol (Fisher, cat. no. A412–500)

Bleaching Solution (see recipe in Reagents and Solutions)

M9 buffer (see recipe in Reagents and Solutions)

10X PBST (see recipe in Reagents and Solutions)

Bovine Serum Albumin (Sigma, cat. no. A9418–5G)

Primary Antibody or Fluorescently Labeled Nanobody/ScFv e.g. K76, anti-PGL-1 (DHSB, Cat# K76, RRID:AB_531836)

Fluorescent Secondary Antibody, e.g. Alex Fluor 488 Goat Anti-Mouse IgG (H+L) secondary antibody (Jackson ImmunoResearch Labs, Cat# 115–545-003, RRID:AB_2338840)

DAPI, 4’,6-Diamidino-2-Phenylindole, Dihydrochloride (Invitrogen, cat. no. D1306)

RNasin^® Ribonuclease Inhibitor (Promega, cat. no. N2111)

Mounting Medium (see recipe in Reagents and Solutions)

VECTASHIELD Mounting Medium (Vector Laboratories, cat. no. H-1000–10)

Liquid Nitrogen

150 mm Petri Dishes (Genesee Scientific, cat. no. 32–106, or equivalent. Other sizes are acceptable.)

Fume Hood (Hamilton Safeaire II, cat. no. 54L2769, or equivalent.)

Stereomicroscope (Nikon, cat. no. SMZ800N or equivalent.)

10 mL Serological Pipets (Genesee Scientific, cat. no. 12–104 or equivalent.)

25 mL Serological Pipets (Genesee Scientific, cat. no. 12–106 or equivalent.)

Pipet Controller (Drummond, cat. no. 4–000-101 or equivalent.)

1000 mL Glass Bottles (VWR, cat. no. 10754–820 or equivalent.)

250 mL Glass Bottles (VWR, cat. no. 89000–236 or equivalent.)

15 mL Centrifuge Tubes (Olympus Plastics, cat. no. 28–103 or equivalent.)

1.5 mL Microcentrifuge Tubes (Olympus Plastics, cat. no. 14–125 or equivalent.)

1.5 mL Amber Microcentrifuge Tubes (Olympus Plastics, cat. no. 22–282AM or equivalent.)

Nutating Mixer (Labnet, cat. no. S0500 or equivalent.)

Centrifuge (Eppendorf, cat. no. 5810 or equivalent.)

Microcentrifuge (Eppendorf, cat. no. 5424 or equivalent.)

Mini Centrifuge (Genesee, cat. no. 27–523 or equivalent.)

Vortex Mixer (VWR, cat. no. 12620–838 or equivalent)

Liquid Nitrogen Container (Thermo Scientific, cat. no. 2122 or equivalent.)

ThermoMixer (Eppendorf, 05–412-503)

8mm 1.5 thickness round cover glass (Electron Microscopy Sciences, cat. no. 72296–08)

Glass microscope slides (VWR, cat. no. 48312–401)

1.5 thickness, 22×22 mm coverglass (VWR, cat. no. 48366–227)

Grace Bio-Lab Press-To-Seal silicon isolator (Sigma, cat. no. GBL664504–25ea)

Widefield Epifluorescent Microscope (DeltaVision Elite or equivalent)

−20 °C Freezer

4 °C Refrigerator

10, 200, and 1000 uL Micropipettes and tips

Protocol steps

1. Perform embryo prep and fixation as in Basic Protocol 1, Steps 1–18.

2. Perform immunofluorescence as in Basic Protocol 1, Steps 19–39, with the following exceptions and additional steps:

   1. Do not perform any smFISH protocol steps.
   2. At step 39, nutate the sample in 1X PBST for 10 min instead of 5.

3. Pellet embryos by centrifuging at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.

4. Pipet or aspirate as much of the supernatant PBST as possible without disrupting the pellet.

5. Counterstain with 1 mL 1X PBST containing 2 uL of 500 ng/mL DAPI for 10 min.

6. Pellet embryos by centrifuging at 2000 × g in 30 sec intervals at room temperature, rotating the tube 180° between each spin until pellet forms.

7. Pipet or aspirate as much of the supernatant PBST as possible without disrupting the pellet.

8. Add 1 mL of 1X PBST directly to sample and nutate for 10 min to wash out excess DAPI.

9. Repeat steps 6–8, followed by steps 6 and 7 (for two 1X PBST washes).

10. Resuspend in 50 uL of mounting medium (or less if the sample is small, see note from Basic Protocol 1, Step 59) and incubate at 4 °C for 30 min to ensure antifade penetrance.

11. Prepare slides as in Basic Protocol 1, Steps 61–66.

See sample Data in Figure 4.

Figure 4. Simplified immunofluorescence in C. elegans embryos.

Immunofluorescence was performed on N2 embryos as described (Basic Protocol 2). Embryos were incubated with 1:20 dilutions of K76 (DHSB, AB_531836) (A) or 1:1000 dilutions 2A4 (DHSB, AB_2618114) (B) primary antibodies, followed by incubation with 1:250 dilutions of Alexa Fluor Goat Anti-Mouse secondary antibody (Jackson ImmunoResearch, AB_2338840, green). In the presence of K76 (anti-PGL-1), P granules are observed (A, top), while 2A4 (anti-ELT-2) stained the intestine-specific ELT-2 transcription factor (B, top). Non-specific binding of the secondary antibody was not observed in either instance (A, B, bottom). Three biological replicates were performed for each experiment. Scale bars represent 10 μm.

BASIC PROTOCOL 3: Single-molecule Fluorescence In Situ Hybridization or single-molecule inexpensive Fluorescence In Situ Hybridization

This protocol describes the workflow for performing smFISH or smiFISH in embryos. It is often desirable to visualize RNA distributions and abundance without the complication of performing immunofluorescence. Performing the smFISH protocol alone simplifies the workflow. Additionally, the fluorescence of some GFP fusion proteins persists through fixation, allowing RNA and protein to be visualized simultaneously without using an immunofluorescence protocol. Moreover, we have also validated the recently developed single-molecule inexpensive Fluorescence In Situ Hybridization (smiFISH) protocol to reduce the cost of visualizing RNA in C. elegans embryos.

Here, users will harvest embryos before fixing and permeabilizing them as in Basic Protocol 1. Primary and secondary smiFISH probes can then be annealed to make low-cost smFISH probe analogs. Embryos are then hybridized with either smFISH or smiFISH probes and placed on slides. The user should then be able to visualize single molecules of RNA in situ as discrete fluorescent puncta.

Materials:

C. elegans strain/genotype of interest (Caenorhabditis Genetics Center, CGC)

Nematode Growth Media Plates (see recipe in Reagents and Solutions)

OP50 E. coli (Caenorhabditis Genetics Center, cat. no. OP50)

100% Reagent Grade Acetone (Fisher, cat. no. A18–500)

100% reagent grade methanol (Fisher, cat. no. A412–500)

Bleaching Solution (see recipe in Reagents and Solutions)

M9 buffer (see recipe in Reagents and Solutions)

DAPI, 4’,6-Diamidino-2-Phenylindole, Dihydrochloride (Invitrogen, cat. no. D1306)

Wash Buffer A (see recipe in Reagents and Solutions)

Wash Buffer B (see recipe in Reagents and Solutions)

Hybridization Buffer (see recipe in Reagents and Solutions)

Mounting Medium (see recipe in Reagents and Solutions)

smFISH probes, 25 uM stocks in 10 mM Tris pH 8.0, e.g. nos-2 probe set (Biosearch Technologies, Custom Probes made using the Stellaris Probe Designer) and/or

Primary smiFISH probes, 100 uM stocks in 10 mM IDTE buffer, e.g. nos-2 probe set (IDT, custom order made using the OLIGOSTAN probe designer)

Secondary fluorescently-labeled smiFISH probes, e.g. 5’ and 3’ Quasar 670 modified FLAP Y probes (Biosearch Technologies, Custom Probes ordered using the custom oligo designer)

RNAse-free Water (Invitrogen, cat. no. AM9922)

VECTASHIELD Mounting Medium (Vector Laboratories, cat. no. H-1000–10)

Liquid Nitrogen

New England Bio Labs Buffer 3 (or 3.1) (NEB cat. no. B7203S)

150 mm Petri Dishes (Genesee Scientific, cat. no. 32–106, or equivalent. Other sizes are acceptable.)

Fume Hood (Hamilton Safeaire II, cat. no. 54L2769, or equivalent.)

Stereomicroscope (Nikon, cat. no. SMZ800N or equivalent.)

10 mL Serological Pipets (Genesee Scientific, cat. no. 12–104 or equivalent.)

25 mL Serological Pipets (Genesee Scientific, cat. no. 12–106 or equivalent.)

Pipet Controller (Drummond, cat. no. 4–000-101 or equivalent.)

1000 mL Glass Bottles (VWR, cat. no. 10754–820 or equivalent.)

250 mL Glass Bottles (VWR, cat. no. 89000–236 or quivalent.)

15 mL Centrifuge Tubes (Olympus Plastics, cat. no. 28–103 or equivalent.)

1.5 mL Microcentrifuge Tubes (Olympus Plastics, cat. no. 14–125 or equivalent.)

1.5 mL Amber Microcentrifuge Tubes (Olympus Plastice, cat. no. 22–282AM or equivalent.)

Nutating Mixer (Labnet, cat. no. S0500 or equivalent.)

Centrifuge (Eppendorf, cat. no. 5810 or equivalent.)

Microcentrifuge (Eppendorf, cat. no. 5424 or equivalent.)

Mini Centrifuge (Genesee, cat. no. 27–523 or equivalent.)

Vortex Mixer (VWR, cat. no. 12620–838 or equivalent)

Liquid Nitrogen Container (Thermo Scientific, cat. no. 2122 or equivalent.)

Thermal Cycler (Bio-Rad, cat. no. T100 or equivalent.)

ThermoMixer (Eppendorf, 05–412-503)

8mm 1.5 thickness round cover glass (Electron Microscopy Sciences, cat. no. 72296–08)

Glass microscope slides (VWR, cat. no. 48312–401)

1.5 thickness, 22×22 mm coverglass (VWR, cat. no. 48366–227)

Grace Bio-Lab Press-To-Seal silicon isolator (Sigma, cat. no. GBL664504–25ea)

Widefield Epifluorescent Microscope (DeltaVision Elite or equivalent)

−20 °C Freezer

4 °C Refrigerator

10, 200, and 1000 uL Micropipettes and tips

Protocol steps:

• 1Perform embryo prep and fixation as in Basic Protocol, Steps 1–18.

Do not perform any immunofluorescence steps. Proceed directly from embryo prep and fixation to the smFISH protocol.

• 2Perform smFISH as in Basic Protocol 1, Steps 41–60, with the following exception at Step 42.

  If using smiFISH probes, the following protocol is required to generate annealed primary + secondary smiFISH probes in place of the simple smFISH probe dilution at step 42.

  1. Combine 8–24 smiFISH primary probes at equimolar ratio and dilute to 0.833 uM in Tris pH 8.0.

     This primary probe mixture is stable at −20 °C indefinitely.

  2. In a PCR tube, prepare a solution of:

     • 2 uL primary probe set

     • 1 uL 50 uM FLAP secondary probe

     • 1 uL NEB Buffer 3 (or 3.1)

     • 6 uL RNAse free water

  3. Anneal primary probe set to fluorophore-labeled secondary probes using the following thermocycling conditions:

     • 1 cycle at 85 °C for 3 min
     • 1 cycle at 65 °C for 3 min
     • 1 cycle at 25 °C for 5 min

     Annealed smiFISH probes are viable at −20 °C for up to at least a week. Treat annealed smiFISH probes as diluted smFISH probes. 2 uL annealed smiFISH probe works well for most hybridizations. smiFISH probes can be used simultaneously with traditional smFISH probes if their fluorophores have distinct spectral profiles. If large fluorescent aggregates are seen when imaging, they may be aggregates of secondary smiFISH probes (See Critical Parameters). If these are observed, an alternate annealing protocol has helped reduce their prevalence in our samples. For this, the smiFISH mixture from Step b is first heated to 95 °C for 3 min before decreasing the temperature by 10 °C increments at 0.1 °C/min to 45°C, incubating for 2 min at each 10 °C interval.
• 3Prepare slides as in Basic Protocol 1, Step 61–66. See sample data in Figure 5.

Figure 5. smFISH and smiFISH in C. elegans embryos.

A. Schematic illustration of smFISH probes. B. Schematic illustration of smiFISH probes. C. nos-2 RNA was visualized using smiFISH (magenta) and smFISH (green). nos-2 smiFISH primary probes used FLAP-Y sequences, and the secondary FLAP-Y probe was 5’ and 3’ dual-conjugated with Quasar 670 fluorophores. nos-2 smFISH probes were 3’ single-conjugated with Cal Fluor 610. D. imb-2 RNA was visualized using smFISH (magenta) and smiFISH (green). imb-2 smFISH probes were 3’ single-conjugated with Quasar 670 fluorophores. imb-2 smiFISH primary probes used FLAP-Y sequences, and the secondary FLAP-Y probe was 5’ and 3’ dual-conjugated with Cal Fluor 610. Embryos were counterstained with DAPI, in blue (C, D). A representative image is shown from three biological replicates, performed using newly annealed smiFISH probes for each replicate. Scale bars represent 10 μm.

REAGENTS AND SOLUTIONS:

Bleaching Solution (per 50 mL):

• 40 mL deionized, distilled water

• 7.2 mL 5 M NaOH (Fisher, cat. no. S318–400)

• 4.5 mL 5% NaHOCl (Ricca, cat. no. 7495.5–32)

• Make fresh and store for no more than one week. Precipitates can form in embryo pellets when using old bleaching solution.

M9 Buffer

• 3 g KH₂PO₄ (Sigma, cat. no. P0662–500G)

• 6 g Na₂HPO₄ (Sigma, cat. no. RDD022–500G)

• 5 g NaCl (Fisher, cat. no. S271–500)

• Add deionized, distilled water (ddH₂O) to 1 L final volume

• Sterilize by autoclaving

• Add 1 mL 1 M MgSO₄ (Millipore cat. no. MX0075–1) using sterile technique after solution cools, to prevent precipitation

• Store indefinitely at room temperature

10X PBST

• 80 g NaCl (Fisher, cat. no. S271–500)

• 2 g KCl (Sigma, cat. no. P3911–500G)

• 14.2 g Na₂HPO₄ (Sigma, cat. no. RDD022–500G)

• 2.4 g KH₂PO₄ (Sigma, cat. no. P0662–500G)

• 1% v/v Tween^® 20 detergent (w/v) (Sigma cat. no. P1379–500ML)

• Add deionized, distilled water to 1 L final volume

• Store indefinitely at room temperature

20X SSC

• 800 mL deionized, distilled water

• 175.2 g NaCl (Fisher, cat. no. S271–500)

• 88.2 g sodium citrate tribasic dihydrate (Sigma, cat. no. S4641–500G)

• pH to 7.0 with 1 M HCl

• Add deionized, distilled water to 1 L and autoclave

• Store indefinitely at room temperature

Wash Buffer A (10% volume/volume formamide)

• 600 uL Stellaris Wash Buffer A (Biosearch Technologies, cat. no. SMF-WA1–60)

• 2.1 mL DEPC-treated RNAse-free water (Invitrogen, cat. no. AM9922)

• Store Wash Buffer A without formamide at 4 °C for up to several months

• When needed, prepare 3 mL for each sample to be hybridized and add 300 uL of deionized formamide (Millipore, cat. no. S4117)

• Prepare Wash Buffer A fresh for each experiment.

Wash Buffer B

• Stellaris Wash Buffer B (Biosearch Technologies, cat. no. SMF-WB1–20 )

• Be sure to add 88 mL RNAse-free water (Invitrogen, cat. no. AM9922) to Wash Buffer B stock before use.

• Store Wash Buffer B with water at 4 °C for up to several months

Hybridization Buffer (10% volume/volume formamide)

• Prepare 110 uL for each sample in an experiment: mix 99 uL Stellaris Hybridization Buffer (Biosearch Technologies, cat. no. SMF-HB1–10/0) with 11 uL of deionized formamide (Millipore, cat. no. S4117). Prepare hybridization buffer fresh for each experiment

• Store hybridization buffer without formamide at 4 °C for up to several months

Mounting Medium (5 mL)

• 2.5 mL 100% glycerol (Sigma, cat. no. G5516–100ML)

• 100 mg N-propyl gallate (Sigma, cat. no. 02370–100G). CAUTION: N-propyl gallate is toxic

• 400 uL 1 M Tris pH 8.0 (Sigma, cat. no. 10708976001)

• Vortex until N-propyl gallate has dissolved

• Store mounting medium in amber tubes or covered in foil at either 4 or −20 °C. The solution is light sensitive

• Store mounting medium until it begins to yellow or crystalize

1 M KPO₄ Buffer pH 6.0

• 108.3 g KH₂PO₄ (Sigma, cat. no. P0662)

• 35.6 g K₂HPO₄ (Sigma, cat. no. P3786)

• Add ddH₂O to 1 L

• The pH should not need to be adjusted, but should be checked

• Autoclave to sterilize

• Store indefinitely at room temperature

Nematode Growth Medium plates

• 3 g NaCl (Fisher, cat. no. S271–1)

• 17 g Agar (Fisher, cat. no. AAJ10654P2)

• 2.5 g Peptone (Fisher, cat. no. BP1420–500)

• Mix in 2 L Erlenmeyer flask

• Add 975 mL ddH₂O

• Autoclave for 50 min

• Add 1 mL of sterile 1 M CaCl₂ (Sigma, cat. no. C4901)

• Add 1 mL of 5 mg/mL cholesterol in 100% ethanol (Sigma, cat. no. C8667)

• Add 1 mL of sterile 1 M MgSO₄ (Millipore cat. no. MX0075–1)

• Add 25 mL of sterile 1 M KPO₄ Buffer

• Swirl to mix

• Add to 150 mm petri dishes using sterile technique

• Once cool and solidifed, seed with 1.5 – 2 mL OP50 E. coli

• Once dry, store for up to a month prior to use

COMMENTARY:

Background Information:

Visualization of proteins and RNAs (Femino et al., 1998; Orjalo et al., 2011; Raj et al., 2008) has been a staple of cell biology for decades. As a result, many methods have been developed for sample preparation, fixation and permeabilization, IF, and smFISH. These methods can be highly divergent in different model systems, or even at different stages of development in a single model organism. In the preceding protocols, we present simplified and optimized methods to fix C. elegans embryos before staining for protein and probing for RNA, in a sequential (Basic Protocol 1), simultaneous (Alternate Protocol 1), or independent (Basic Protocol 2 and Basic Protocol 3) manner.

In situ visualization of biological molecules requires that samples be preserved to mirror living cells as closely as possible. Modern fixation was developed in the sixteenth century primarily to preserve tissue samples for histological studies, first using solutions of alcohols, waxes, acetic acid, and arsenic (Cole, 1951). As the mechanisms of fixation were characterized, researchers identified countless reagents for preserving samples using diverse mechanisms. Common fixatives include formaldehyde/formalin, picric acid, or organic solvents such as methanol, ethanol, and acetone. Formaldehyde/formalin acts by creating crosslinked, covalent chemical bonds in the sample, primarily at lysine residues. Notably, formalin can also cause C-T and G-A mutations on DNA sequences, as characterized by PCR (Williams et al., 1999). Moreover, formaldehyde/formalin-fixation affects tertiary amines in RNA sequences, resulting in modification of up to nearly 40 % of As and Cs in formalin-fixed tissues (Masuda et al., 1999). Due to the high degree of alteration that occurs on nucleic acids, formaldehyde/formalin-fixation is not an ideal fixative for nucleic acid visualization. Similarly, picric acid is a problematic fixative for smFISH experiments, as its low pH catalyzes nucleic acid degradation and leads to poor hybridization (Bonin et al., 2005; Tyrrell et al., 1995). As an alternative to crosslinking-fixatives, alcohols and other organic solvents have been identified as superior nucleic acid fixatives (Srinivasan et al., 2002). Alcohols and organic solvents, such as ethanol, methanol, and acetone, function by dehydrating clathrate water molecules around proteins and nucleic acids, thus precipitating biological molecules into a fixed state, without significant chemical alteration. As with crosslinking fixatives, however, alcohols and organic solvents have their detriments. These fixatives can disrupt cell membrane structures, cytoplasmic organelles, and soluble cell components such as microtubules (Hoetelmans et al., 2001; Vielkind & Swierenga, 1989). However, due to their preservation of nucleic acid composition, alcohols and organic solvents are ideal fixatives for single-molecule RNA detection assays. Further, we have found that liquid nitrogen freezing combined with short fixations alcohol/organic solvent fixation allows efficient antibody penetration through the embryonic permeability barrier, and without disrupting the protein epitopes we have targeted through IF, as some previous studies have shown (Levitt & King, 1987). Ultimately, the use of methanol and acetone fixation greatly simplifies slide preparation for IF samples by removing the need to mechanically permeabilize the embryos on a slide while circumventing many of the challenges that standard fixation protocols using formaldehyde/formalin and picric acid pose. By permitting an in-tube fixation and permeabilization compatible with protein and RNA detection, samples fixed in this manner will be ready for both IF and smFISH protocols.

Immunofluorescence is a standard protocol in cell biology for the visualization of proteins in fixed cells. By targeting a protein of interest with fluorescently labeled primary/secondary antibody pairs, the distribution, abundance, timing, and other features of protein expression can be visualized (J. Duerr, 2006). Since the adoption of C. elegans as a model organism, various adaptations have been implemented to overcome the specific challenges of visualizing protein in this organism. These adaptations provide information for antigen production, peptide coupling, antibody purification, fixation conditions, and protocols related to IF in C. elegans (J. Duerr, 2006; Strome & Wood, 1982; Ward & Klass, 1982). However, most have focused on larval stages of development and are not optimized for embryos. Most protocols use some combination of reducing reagents, enzymatic treatments, formaldehyde fixation, and “Freeze-Cracking” mechanical disruption. The standard immunofluorescence freeze-cracking procedure is the compression of samples between slides, followed by their rapid separation, not to be confused with freeze-cracking of the eggshell in liquid nitrogen (J. S. Duerr, 2013). The single-tube fixation protocol described here mitigates some issues with traditional IF protocols by removing reducing reagents and enzymatic treatments while permeabilizing the eggshell, thereby simplifying the entire protocol, from fixation to slide preparation. This adapted IF protocol stains proteins of interest while preserving the RNA content of the embryo. Users can then visualize the protein immediately or continue to perform an smFISH protocol subsequently (Strome & Wood, 1982)(Wiesenfahrt et al., 2015).

The current gold standard for in situ single-molecule RNA detection is single-molecule Fluorescence in situ hybridization (smFISH). smFISH allows researchers to characterize the developmental dynamics of RNA expression, including abundance, localization, decay, and the onset of transcription. RNA In Situ Hybridization was initially developed using radiolabeled and enzymatically-driven colorimetric assays with low spatial resolution. It then evolved to use antisense probes conjugated to multimeric fluorophores, leading to high background signal and low specificity (Young et al., 2020). In current smFISH protocols, single-molecule RNA visualization occurs by annealing a series of ∼24–48 fluorescently-labeled short antisense oligonucleotide probes to a transcript of interest in fixed cells/animals (Femino et al., 1998; Orjalo et al., 2011; Raj et al., 2008). Annealing multiple fluorescent probes to an RNA produces a discrete, punctate signal for individual RNA molecules in situ. smFISH probes can be designed and synthesized in the lab (Ji & Oudenaarden, 2012; Raj et al., 2008) or ordered as a set from Biosearch Technologies (Novato, CA). Some typical fluorophores include Cy5, Quasar 670, Alexa 594, Cal Fluor 610, and Fluorescein, among many others. In RNA FISH experiments, it is crucial to obtain the highest possible signal-to-noise ratio (SNR) to ensure reliable interpretation of the data.

One common question surrounding smFISH is whether commercial reagents (i.e., Stellaris) are superior to homemade ones (Ji & Oudenaarden, 2012; Orjalo et al., 2011). By comparing the signal-to-noise ratio of transcripts imaged by smFISH, we have found Stellaris buffers, in general, to perform better than homemade buffers (for example, see FIGURE 6). Another common concern with smFISH experiments is photolability. Due to the relatively low signal, high laser powers, and the small number of fluorophores (~24–48) utilized in smFISH experiments, photobleaching can occur rapidly. Photobleaching is of particular concern with thick samples that must be imaged through many Z stacks, as is the case with C. elegans embryos (~12–20 um thickness as prepared in Basic Protocol 1, or ~60–100 stacks per embryo at 0.2 um spacing between Z-stacks). One of the primary causes of photobleaching is the degradation of fluorophore molecules by oxygen radicals produced upon laser excitation (Greenbaum et al., 2000). Therefore, free-radical scavenging antifade solutions are commonly used to reduce the degree of experimentally-induced photobleaching. We have observed that the optimal antifade solution can vary depending on the probe set or fluorophore (FIGURE 7). In our hands, VECTASHIELD, N-propyl gallate, or a mixture of the two, provide the best signal stability for Cal Fluor 610 and Quasar 670 labeled RNAs in C. elegans embryos. We generally avoid probes conjugated with fluorescein, as they tend to have very low signal-to-noise ratios.

Figure 6. Stellaris buffers provide higher signal-to-noise ratios than homebrew buffers.

Signal-to-noise ratios were calculated for each RNA puncta identified when smFISH was performed using homebrew (red) or commercial Stellaris (blue) buffers. The signal-to-noise ratio was calculated by identifying RNA spots using FISHquant (Mueller et al., 2013) before using the ImJoy SNR plugin (Imbert et al., 2021). In short, the SNR plugin compares the intensity at the coordinates of RNA puncta identified by FISHquant to the average intensity of a sphere surrounding the spot to calculate SNR. Four Stellaris smFISH probe sets were used, erm-1 conjugated to Cal Fluor 610, imb-2 conjugated to Quasar 670, nos-2 conjugated to Quasar 670, and set-3 conjugated to Cal Fluor 610. Individual dots represent the average SNR in one embryo. Three biological replicates were performed for each experiment, and 15 embryos were quantified for each condition over the three replicates. P values from Benjamini-Hochberg corrected t-tests are shown (0.05 > *, 0.005 > **, 0.0005 > ***).

Figure 7. Effect of antifade composition on smFISH signal intensity.

The mean fluorescence intensity of smFISH signal over 100 exposures was measured in embryos using various antifades and their combinations. Experiments were performed using four different smFISH probe sets: erm-1 conjugated to Cal Fluor 610, imb-2 conjugated to Quasar 670, nos-2 conjugated to Quasar 670, and set-3 conjugated to Cal Fluor 610). A. Representative images of the first and final acquisitions for imb-2 (top) and erm-1 (bottom) RNAs using VECTASHIELD and N-propyl gallate (left), VECTASHIELD only (middle), and ProLong Diamond (right) antifades. B. The average mean intensity throughout imaging was normalized to the intensity of first acquisition for each embryo. The shaded region represents the standard error of the mean for each exposure. Three biological replicates were performed for each experiment, and no less than nine embryos were quantified for each condition.

Because each probe in a standard smFISH probe set requires chemical conjugation with fluorophores and is specific to each transcript, smFISH probe sets are relatively expensive (Femino et al., 1998; Orjalo et al., 2011; Tsanov et al., 2016). Targeting a single RNA typically costs in the range of ~$500. Recently, Tsanov et al. outlined a straightforward, flexible method for reducing the cost of single-molecule RNA detection: single-molecule inexpensive Fluorescence In Situ Hybridization (smiFISH). smiFISH brings down the cost of single-molecule RNA detection by taking advantage of a single, universal fluorophore-labeled secondary probe annealed in vitro to gene-specific primary probes (Figure 5A). Primary smiFISH probes contain two main parts facilitating efficacy and cost reduction: the gene-specific region complementary to the transcript of interest and the FLAP region complementary to the fluorescently labeled secondary probe. In situ, the complementary region of the primary probes binds to the target RNA, while its FLAP region is annealed to a fluorophore-labeled secondary FLAP probe. This regime significantly reduces the cost of single-molecule RNA visualization by eliminating the need to create chemically conjugated probe sets for each specific target RNA. smiFISH primary probes can be designed as described using the R script Oligostan (Tsanov et al., 2016). Primary probes can be ordered in 96-well plates from IDT on the 25 nmol scale prediluted to 100 uM in IDTE buffer pH 8.0. Alternatively, if ordering 96 or more individual probes, oligos can be ordered on the 500 pm scale, providing ample primary probes for hundreds of experiments. For most experiments, ~12–16 primary probes per transcript is sufficient, although testing as few as 8 primary probes has produced discernable single-molecule spots in C. elegans embryos (Parker et al., 2020). An increased number of primary probes typically increases the signal-to-noise ratio for any given transcript. Secondary FLAP probes can also be ordered as 5’ and/or 3’ single- or dual-fluorophore-labeled oligos from either Biosearch Technologies or IDT (Coralville, Iowa). We have previously demonstrated that smiFISH performs as well as traditional smFISH in C. elegans embryos (Parker et al., 2020). We found that smiFISH faithfully reproduces the sensitivity, spatial resolution, and reliability of smFISH probes in embryos. Notably, smiFISH is less effective in larval stages than smFISH using our protocols, possibly due to lower larval permeability, preventing smiFISH probe entry. Here, we provide an example, comparing nos-2 or imb-2 smFISH and smiFISH probes on the same sample (FIGURE 5). Using the smFISH or smiFISH protocols presented here facilitates visualization of single RNA molecules in fixed samples. Additionally, if the smFISH protocol is performed after IF, as described in Basic Protocol 1, both protein and RNA can be imaged in the same sample.

Traditional approaches to visualizing mRNA and protein simultaneously in C. elegans have relied on the visibility of a fluorescently-tagged protein persisting under RNA labeling conditions. However, these fluorescent protein tags can often bleach during fixation. The combination of IF and smFISH protocols facilitates the simultaneous visualization of protein and RNA without requiring the genetic manipulation of fluorescently tagging a protein. Yoon et al. developed a protocol to perform IF and smFISH concurrently in C. elegans, but it requires manual dissection of animals to isolate the antibody-permeable gonad (Yoon et al., 2016). We provide a protocol that combines both IF and smFISH in C. elegans embryos with a single-tube approach, requiring no genetic manipulations and using techniques accessible to novices. These protocols permit a broader range of C. elegans experimentalists to compare protein and RNA distributions in the same sample.

CRITICAL PARAMETERS:

Permeabilization and fixation:

C. elegans embryos are well isolated from environmental contaminants. This is partly due to the permeability barrier, a membranous barrier that prevents fluid exchange between the embryo and the environment (Olson et al., 2012). The fixative reagent and incubation time are important for permeabilizing the embryo to antibodies, which are roughly 20X the mass and radius of smFISH probes (Ab ~ 150 kDa and ~ 60 A, 20mer oligo ~ 7.5 kDa and ~ 3 A (Fernandes et al., 2002; Hawe et al., 2011).). In our experience, a brief methanol fixation, and liquid nitrogen freeze cracking followed by a quick acetone fixation, was most effective at allowing antibodies to pass through the eggshell and permeability barrier, while maintaining antigen recognition and FISH probe accessibility. We found that the use of acetone was necessary for antibody staining. We interpret this result as acetone solubilizing permeability barrier components, thus increasing the size of molecules that can enter the embryo, although we have not rigorously examined the effective pore size under different fixation conditions. Our experiments with longer fixation times with both methanol and acetone reduced antigen recognition by antibodies (as well as GFP fluorescence for protein fusions). Moreover, the use of formalin/formaldehyde reduces the binding and photostability of FISH probes. Some antigens are likely more compatible with different fixatives, and, Duerr (J. Duerr, 2006) describes alternative fixation strategies if the fixation conditions presented here are incompatible with an antigen of interest. If alternative fixation strategies must be pursued, it is crucial to keep in mind their effect on the permeability of the eggshell and permeability barrier.

Sequential IF/FISH protocol.

Simultaneous detection of an RNA and its cognate protein can reveal a wealth of information regarding the expression patterns, regulation, and functions of genes. However, the combination of IF and FISH is often challenging due to slight incompatibilities in traditional protocols. When immunofluorescence is performed in series with smFISH, all reagents must be RNAse-free, where possible. It is absolutely essential that solutions that contain BSA be treated with an RNAse inhibitor to prevent RNA degradation. If RNAse-free reagents (not including BSA) are not available or if degradation is occurring, the addition of RNAse inhibitors can slow RNA degradation. Though most reagents listed do not cause apparent RNA degradation, we typically try to only use RNAse inhibitor in steps containing BSA because it is expensive to include with every reagent.

Simultaneous IF/FISH protocol.

A simplified protocol can often be utilized if performing IF with a high-affinity nanobody or single-chain variable fragments (ScFvs). Under these circumstances, simultaneous smFISH/IF can be achieved by performing the smFISH protocol (Basic Protocol 3) with the simple addition of fluorescently labeled nanobody or ScFv to the hybridization buffer before overnight incubation with the FISH probes and sample. It is unclear, however, why some nanobodies and ScFv work with this simplified protocol. Perhaps their smaller size allows better permeation during hybridization, or their higher affinity is more compatible with RNA FISH hybridization temperatures.

Validation of antibodies:

It is essential to validate antibody function and specificity in any IF experiment. Primary antibodies can be validated using null strains or RNAi to ensure that the antibody is binding specifically to the target antigen. Secondary antibodies can be tested for specificity by incubating them in the absence of primary antibodies to ensure no staining of endogenous antigens. Should an antibody have some non-specific binding, it may be possible to increase specificity by depleting the antibody using a null allele (J. Duerr, 2006). It is also necessary to optimize antibody concentrations over one or two orders of magnitude (starting with manufacturer recommendations) to determine optimum conditions for antigen detection without non-specific binding. Note that optimum antibody concentrations can change with experimental conditions where protein concentrations vary significantly. Because altering antibody concentrations can make downstream quantification inaccurate, it is beneficial to use identical staining conditions when possible.

Validating new smFISH/smiFISH probe sets:

There are several ways to validate new smFISH and smiFISH probe sets for target specificity and labeling efficiency. Testing a probe set in a wildtype and deletion strain for the target of interest ensures the probe set is specifically binding only when the RNA is present. If a deletion allele is not available, RNAi can be utilized to a similar end. However, it is important to note that residual fluorescent signal may be present after RNAi due to incomplete knockdown or partial degradation of the targets. Target specificity can also be determined by targeting a transcript with two separate probe sets in different colors, which should colocalize if the probes are specific. The labeling efficiency of a probe set can be determined by comparing transcript abundance found using smFISH data to other sources, such as RT-qPCR, digital-droplet PCR, or quantitative sequencing data.

Positive controls:

Positive control smFISH probe sets should be consistently employed. These probe sets have the added benefit of marking specific cell lineages or developmental stages, and thereby identify the embryo’s orientation or stage. By comparing the performance across replicates, researchers can identify outliers or problems in protocol execution. When troubleshooting, smFISH probe sets that anneal to highly abundant RNAs (polyA) or previously validated targets can ensure success and aid in diagnosing issues.

Low Signal to Noise Ratio (SNR):

Since C. elegans embryos are relatively thick (~20–30 um), the use of widefield microscopy will capture out-of-focus signal from non-focal Z-planes, thereby decreasing the SNR. Embryos can be flattened during slide preparation to improve SNR. We have found that samples from ~12–20 um thick have an optimal SNR without perturbing sample morphology. While pressing down on embryos does not seem to affect their morphology, any lateral motion during slide preparation will shear embryos, so pressing directly down when making slides is essential.

Crosstalk of smiFISH secondary probes:

It is possible to use two independent primary probe sets containing the same FLAP sequence without crosstalk. Tsanov et al. demonstrated that multiple primary probe sets containing the same FLAP sequence could be utilized in the same sample without observable mislabeling of their target transcripts by annealing them to secondary probes labeled with distinct fluorophores (i.e., probeset-1 FLAP-Y-Cal Fluor 610, probeset-2 FLAP-Y-Quasar 670). We have previously confirmed this in the C. elegans embryo (Parker et al., 2020).

TROUBLESHOOTING:

Please see Table 1 for a list of common problems with the protocols, their causes, and potential solutions.

Table 1.

Troubleshooting

Problem	Possible Cause	Solution
No antibody staining	Antibody did not bind its target	Use validated primary and secondary antibody pair as positive control
	Overfixed sample	Do not fix for more than for a total of 10 min between methanol and acetone
	Poor permeabilization	Check embryo permeability with antibody sized fluorescent dextran (~150 kDa)
	Wrong concentration of antibody	Titrate antibodies
	Primary antibody doesn’t bind	Validate primary antibody using a previously validated secondary antibody
	Secondary antibody doesn’t bind	Validate secondary antibody using a previously validated primary antibody
No antibody staining (Alternate Protocol 1 only)	Antibody is not compatible with FISH buffers	Use Basic Protocol 1 or 2
Protein or RNA looks abnormal	Sample was damaged	Use an RNA or protein marker to ensure cell morphology is in tact
No RNA spots	smFISH probe did not bind its target	Use validated smFISH probes as a positive control
	RNA was degraded	Ensure reagents are RNAse-free or use RNAse inhibitor at all steps
No protein/RNA spots	Sample was stored too long	Reduce length of incubations during IF and/or smFISH steps
Rapid RNA spot photobleaching	Wrong antifade mixture	Check antifade specs to make sure they are compatible with the FISH fluorophore
Rapid RNA spot photobleaching	Antifade hasn’t permeated the embryo	Give the embryos time to incubate after slide preparation to ensure the antifade penetrates the sample
Rapid RNA spot photobleaching	Not imaging from high wavelength to low wavelength	Image the entire z-volume using the highest wavelength before moving to the next highest and so on
Low SNR	Sample is too thick	Press more firmly when preparing slides, aiming for a thickness of ~12–14 um
	Low number of probes	Increase number of probes
Low SNR (smiFISH only)	Single fluorophore-labeled secondary probes	Order dual 5’ and 3’ labeled secondary smiFISH probes
Impossible to design more than 8 probes for target transcript	Short transcript	Consider amplification-based RNA FISH methods ((Choi et al., 2016, 2018; Marras et al., 2019; Wang et al., 2012; Xia et al., 2019))
Bright aggregates when using smiFISH	Secondary probe aggregation	Vortex aggressively smiFISH probes after annealing and pellet aggregates using minifuge spin and/or use the alternate slow annealing program
Low embryo yield (Basic Protocol 1)	No detergent in PBS	Use PBST. Embryos will stick to the tube without it.
Clumpy embryos (Basic Protocol 1)	Undetermined	Vortex aggressively during all IF steps and break up clumps with pipette when preparing slides

UNDERSTANDING RESULTS:

Performing the RNA FISH or immunofluorescence protocols described in this article should result in dual-labeled, stacked images reporting fluorescently-stained protein targets and clear punctate spots for RNA targets either in combination or independently (Figures 2–5). For instance, in Figure 2A and 2B, our sequential immunofluorescence and smFISH protocol demonstrates that the PGL-1 antibody stains large fluorescent condensates known as P granules, while probing for nos-2 and cpg-2 RNAs showed overlapping clusters of these RNAs localized within P granules. Overall, these results demonstrate a robust series of protocols for producing high-quality data examining the distributions of proteins and RNAs of interest in the C. elegans embryo.

smFISH and smiFISH data analysis:

There are several routes for interpreting smFISH data, depending on the biological questions at hand. These analyses range from characterizing the quality of the data, counting the number of RNAs in the samples, or identifying spatial distributions of RNA within cells of interest.

The most common method for quantification of smFISH data is counting the number of RNAs within the sample. For this purpose, some commonly used tools are FISH-quant (Mueller et al., 2013) and StarSearch (Raj et al., 2008). These algorithms function by enhancing spot signals through various filtering methods, setting a threshold for RNA spot detection, and identifying individual spots. Thresholds are often set manually by testing a range of intensity values. Therefore, when plotting these values against the number of detected spots, a plateau can often be seen corresponding to threshold values separating RNA spots from lower intensity noise. When performing spot detection analysis of smFISH data, it is imperative to ensure that the SNR of the data is sufficient to identify spots unambiguously. SNR can be calculated using an ImJoy plugin, which compares the intensity of a detected spot to the surrounding background intensities (https://github.com/fish-quant). In our experience, if SNR values are below ~3–4, spot detection becomes less reliable. When analyzing smFISH data using FISH-quant or StarSearch, if there is no apparent plateau of RNA counts over various threshold values, the SNR is likely too low for accurate RNA spot detection.

As smFISH has become more widely utilized, novel methods of analysis beyond spot counting are continuously being developed. For instance, FISH-quant has been ported from Matlab to an open-source implementation in Python and successfully applied to two large-scale screen projects (Chouaib et al., 2020; Safieddine et al., 2021). This package includes methods for detecting RNAs, deconvolving overlapping RNAs to increase the counting accuracy of highly abundant or clustered RNAs (Chouaib et al., 2020; Parker et al., 2020), measuring the signal-to-noise ratio of an image (https://github.com/fish-quant), and identifying diverse subcellular localization patterns of RNA (Chouaib et al., 2020; Samacoits et al., 2018). Further, to facilitate its usage by non-specialists, several plugins providing user interfaces for the data analysis platform ImJoy (Ouyang et al., 2019) were developed. As more labs adopt smFISH methodologies and more high-throughput methods of in situ RNA detection develop (Eng et al., 2019; Lubeck et al., 2014; Moffitt et al., 2016; Querido et al., 2017; Xia et al., 2019), more sophisticated analysis methods are likely to arise. An exciting initiative is Starfish, an open-source software suite with the goal of building a unified data-analysis tool and file format for several spatial transcriptomic techniques (Perkel, 2019).

IF data analysis:

Standard methods of analysis for IF experiments include measuring the total internal fluorescence and measuring colocalization between different markers. These methods require imaging conditions, such as laser intensity and exposure times, to be held constant across samples and replicates. We will highlight publicly available tools for analysis here; however, most microscopes ship with instrument-specific software packages capable of performing these analyses. Total internal fluorescence compares the intensity of a protein visualized by IF in a control sample and an experimental condition, such as an RNAi knockdown or protein knockout. Total internal fluorescence can be measured over the total volume of the embryo, or, if specific regions must be analyzed, regions of interest can be masked automatically or manually. Regardless of whether particular segmentations are required, these analyses can be performed relatively quickly in FIJI Is Just ImageJ (FIJI) (Schindelin et al., 2012; Schneider et al., 2012). Additionally, FIJI plugins are available to analyze a protein of interest’s colocalization with another fluorescent marker. When performing colocalization analyses, it is crucial to consider optimal uses for any given colocalization metric, as there are well-documented circumstances where these metrics can be misleading (Dunn et al., 2011). Helpful instructions for segmentation, colocalization analysis, and much more can be found at https://imagej.net/.

Combined IF/FISH data analysis:

As with the IF data analysis, colocalization analyses may be performed on combined IF/FISH data. However, due to the punctate nature of FISH signals, RNA spots may not overlap with a colocalization marker as well as expected, resulting in deceptively low colocalization coefficients. This can occur for several reasons. First, the small total volume of RNA puncta can lead to high variability in colocalization. This variability is compounded by the low temporal resolution of fixed cell experiments and the stochastic movements of RNA in the cell, even for tightly localized transcripts. For these reasons, several groups are developing novel metrics for comparing RNA and protein data and analyzing the spatial relationships between them. For instance, by spatially modeling the coordinates of each RNA puncta and comparing their distributions to other RNAs or organelles, it is possible to identify RNA patterning at various cellular features such as cortical membranes, nuclear membranes, condensates/puncta, cellular protrusions, centrosomes, and more (Chouaib et al., 2020; Parker et al., 2020; Safieddine et al., 2021; Samacoits et al., 2018).

TIME CONSIDERATIONS:

The entire procedure described in Basic Protocol 1 can be completed in 2–3 days. On the first day, embryos are harvested from gravid worms, fixed, and stained overnight with primary antibodies. On the second day, fluorescently labeled secondary antibodies are added to complete the immunofluorescence protocol before performing smFISH. smFISH probes can then be incubated for as few as 4 hours before washing; however, overnight smFISH probe hybridization vastly improves the RNA signal. Thus, smFISH probes are washed from the sample on the evening of the second day or the beginning of the third day before imaging. Alternate Protocol 1, Basic Protocol 2, and Basic Protocol 3 can be performed in 2 days. Once again, the embryos are harvested on the first day before overnight incubation with both antibody and smFISH probes (Alternate Protocol 1), primary antibody alone (Basic Protocol 2), or smFISH/smiFISH probes alone (Basic Protocol 3). In both Alternate Protocol 1 and Basic Protocol 3, unbound antibodies/probes are washed on the second day before imaging. In Basic Protocol 2, immunofluorescence is first completed by adding secondary antibodies prior to washing and imaging.

DATA AVAILABILITY STATEMENT:

The data that support the findings of this study are openly available at the Colorado State University repository at http://dx.doi.org/10.25675/10217/233940.