Information

What do the acronyms in C.elegans neuron names stand for?


In this site, I see a variety of acronymic names for C.elegans neurons but what do these names mean (for example AVAL, AVAR)?


Unfortunately, there doesn't seem to be a very systematic nomenclature. Some of the letters are acronyms while others describe a class of neurons. The letters V and D usually describe the ventral and dorsal counterpart of a pair of neurons. The same goes for L and R (left and right).

This list names all C. elegans neurons and you can see that, for example, the "ring interneurons" or motor neurons start with RI or RM. "Inner labial" or "outer labial" neurons are called IL or OL. Ventral cord motor neurons start with a V and their different classes are called VA, VB, VC and VD plus a number. The classes are determined by lineage origins or historic reasons ;) And there are a lot of exceptions to these rules, because their historic names remain even though we continue to learn more about these neurons.

The example AVAL/AVAR are ventral cord interneurons. They form a class with other ventral cord interneurons called AVB, AVD, AVE. So, the first two letters refer to the main class, the third letter to the subclass. The L or R refer to the left and right member of this pair.

Some friends working in C. elegans neuroscience told me that you basically just treat these letter codes as individual names for the neurons without bothering about their origin and meaning too much.


The hourglass organization of the Caenorhabditis elegans connectome

Affiliations Department of Complex Trait Genetics, Center for Neurogenomics and Cognitive Research, Amsterdam Neuroscience, Vrije Universiteit Amsterdam, Amsterdam, The Netherlands, Department of Clinical Genetics, Amsterdam University Medical Center, Amsterdam Neuroscience, Amsterdam, The Netherlands

Roles Conceptualization, Methodology, Supervision, Writing – original draft

Affiliation School of Computer Science, Georgia Institute of Technology, Atlanta, Geogria, United States of America


INTRODUCTION

Unlike nonliving matter, living beings have developed various mechanisms to sense and respond to the fluctuations of environmental conditions, especially when these become highly unfavorable (usually designated as stress conditions). Entering a resting dormant state will allow some organisms to effectively withstand the adverse conditions and, thus, has been adopted as a common survival strategy by both prokaryotes (e.g., E. coli and M. tuberculosis) and eukaryotes (e.g., yeasts, ciliates, and nematodes).

Many animals have long been noticed to enter an inactive state, known as hibernation, during the winter when food availability is limited. The fertilized eggs of soil nematode C. elegans would normally develop into their adult and reproductively mature stage within 3–5 days, after quickly going through the L1 to L4 larval stages, molting at the end of each stage. Nevertheless, C. elegans is also able to enter a quiescent state, designated as dauer (a German word meaning “enduring or persisting”) stage, usually at the end of the L2 (or less commonly of the L1) larva stage, under harsh conditions such as lack of food, high-population density, or temperature increase (see Fig. 1) ( 2 , 3 ). The dauer state allows C. elegans to commonly survive for up to 120 days (instead of the normal 20 days!), as well as to effectively disperse via phoresy (relationship in which one organism transports another organism of a different species) to a more favorable condition place.

The life cycle of C. elegans. Dauer is an alternative larval stage, taken when the environmental conditions are unfavorable for further growth. Dauer larvae appear to be nonaging and can live nearly 10 times their normal lifespan (about 20 days). The length of time the animal spends at a certain stage is indicated.

The dauer larvae are easily distinguished from other developmental stages by presenting an adverse-resisting and energy-saving state: the animals stop taking any food, appear thin and dense, usually remain motionless, but can remarkably respond to some stimuli. Also, unfed dauer larvae possess a dauer-specific behavior known as nictation, in which a larva mounts a projection and stands on its tail, waving its head in the air. The dauer state is also correlated with a developmental suspension of the reproductive organs. On returning to favorable living conditions, the larva exits the dauer state and resumes the normal development process by first entering the L4 larva stage. Genetic studies have been quite effective in identifying genes whose mutation either promotes or suppresses the dauer formation. However, biochemical studies to understand how encoded proteins are actually involved in the dauer formation process and eventually generate the dramatic morphological and metabolic transformations still remain scarce. This review emphasizes on the progresses achieved so far in understanding the biochemical process of dauer formation.


2. C. elegans basics

2.1. Growth and maintenance

C. elegans , although often mischaracterized as a soil nematode, can most easily be isolated from rotting vegetable matter, which contains an ample supply of their bacterial food source (Barrière and Félix 2014). In the laboratory, animals are normally grown on agar plates containing a lawn of the bacterium Escherichia coli . Once the animals deplete the bacteria, they utilize their fat supply. Without food, the development of young larval stage animals is arrested. As a result of entering this stasis, animals can survive for at least a month (often starved plates can be usefully kept for up to six months at 15°), and as stocks, they do not require constant feeding. Whenever healthy, growing animals are needed, a piece of the agar from the old plate can be transferred to a new plate with bacteria. The animals move to the new bacteria and resume their development.

Several other features greatly facilitate the maintenance of C. elegans stocks and their experimental use. First, because C. elegans is a self-fertilizing hermaphrodite (see the following section), a single animal can populate a plate. Second, animal populations can be frozen for years and revived when needed. Third, the animal's small size means that many can be grown in a small space. Fourth, animals can be grown at temperatures ranging from 12° to 25° their Q10 for growth is

2 (that is, an increase of 10° speeds up growth twofold). Growth at different temperatures makes it possible to control the rate of animal development and assists in the isolation and use of temperature-sensitive mutants. Continual growth above 25° is not possible because the animals become sterile. The upper temperature limit can be a problem if animals are kept on bench tops (instead of temperature-controlled incubators) in rooms that are too warm. Shorter exposures to higher temperatures are possible for heat shock experiments and to increase production of males (Sulston and Hodgkin 1998). Fifth, animals can be synchronized by isolating newly hatched larvae or by treating gravid adults with bleach (which decontaminates by killing everything but embryos) and isolating eggs, which are resistant to bleach treatment. Sixth, to facilitate biochemical studies animals can be grown in bulk in liquid medium. ”Worm sorters” such as the COPAS Biosorter are also available to quickly select large quantities of individual worms with desired characteristics. Finally, one does not need especially expensive equipment beyond a good dissecting microscope and a compound microscope to work with this animal. Overall, the animals are inexpensive and convenient to maintain.

2.2. Sexual forms and their importance

Wild-type C. elegans has two sexual forms: self-fertilizing hermaphrodites and males (Figure 2 and Figure 3, A and B). The gonad of hermaphrodites forms an ovotestis that first produces haploid amoeboid sperm that are stored in the spermatheca in the L4 stage and then near adulthood the germ line switches fate to produce much larger oocytes. Essentially hermaphrodites are females whose gonads temporarily produce sperm before they produce oocytes. Hermaphrodites can produce up to 300 self-progeny that are fertilized by the stored sperm. If mated with males, hermaphrodites are capable of producing

1000 offspring, indicating that hermaphrodite-produced sperm is a limiting factor in self-fertilization. Both sexes are diploid for the five autosomal chromosomes. The sexes differ in that hermaphrodites have two X chromosomes and males have a single X chromosome— C. elegans has no Y chromosome—and the genotype of males is referred to as XO. Sex is determined by the X to autosome (X:A) ratio (Zarkower 2006). The majority of offspring produced by self-fertilization are hermaphrodites only 0.1-0.2% of the progeny are males due to rare meiotic non-disjunction of the X chromosome. Because hermaphrodites make their own sperm, in genetic crosses self-progeny (oocytes fertilized by the hermaphrodite's sperm) need to be distinguished from cross-progeny. For example, when hermaphrodites homozygous for a recessive mutation causing a visible mutant phenotype are mated to wild-type males, the self-progeny hermaphrodites show the mutant phenotype and the cross-progeny hermaphrodites do not.

Figure 3. C. elegans anatomy. Major anatomical features of a hermaphrodite (A) and male (B) viewed laterally. (A) The dorsal nerve cord (DNC) and ventral nerve cord (VNC) run along the entire length of the animal from the nerve ring. Two of the four quadrants of body wall muscles are shown. (B) The nervous system and muscles are omitted in this view, more clearly revealing the pharynx and intestine. (C) Cross-section through the anterior region of the C. elegans hermaphrodite (location marked with a black line in A) showing the four muscle quadrants surrounded by the epidermis and cuticle with the intestine and gonad housed within the pseudocoelomic cavity. Images modified from those found at www.wormatlas.org (Altun et al . 2002-2015).

Self-fertilizing hermaphrodites provide several advantages for genetic analysis. First, self- fertilization (often referred to as selfing) simplifies maintaining stocks because a single animal can give rise to an entire population. Second, as Sydney Brenner (1974) has written, “the animals are driven to homozygosity,” i.e., populations of hermaphrodites tend to lose heterozygotes (because hermaphrodites cannot mate with other hermaphrodites). Thus, strains that are mutagenized are essentially isogenic. Third, selfing follows the standard Mendelian rules of segregation, so a parent that is heterozygous for a recessive trait will produce the standard 1:2:1 pattern of segregation, such that 25% of the progeny will be homozygous for the mutant allele and display the autosomal recessive trait. Thus, selfing reduces greatly the effort needed to find such mutants. Fourth, mutants with neuromuscular defects that impair the ability to mate can still be maintained in the laboratory. In fact, only eleven of the 302 nerve cells of the hermaphrodite: eight ADF, ASG, ASI, and ASJ neurons, which when killed as a group cause the animals to become dauer larvae (Bargmann and Horvitz 1991), the two CAN cells (Forrester and Garriga 1997), and the M4 neuron in the pharynx (Avery and Horvitz 1989) are known to be essential to support development to the reproductive adult stage. Fifth, the viability of even severely defective mutants and their ability to self-fertilize allows for easy screens for modifier (enhancer and suppressor) mutations. Such screens have been exceptionally useful and informative. For example, lin-12 mutants ( C. elegans nomenclature is outlined in Box 1) are defective in vulval development and components of the LIN-12/Notch signaling pathway have been identified with both suppressor and enhancer screens (Greenwald and Kovall 2013).

Box 1. C. elegans Nomenclature

Genetic nomenclature differs from species to species. Here, we describe the major terms used in C. elegans research. A more complete description can be found at http://www.wormbase.org/about/userguide/nomenclature.
Nomenclature at a glance 1 : Definition
ZK154.3 Systematic gene identification (3 rd ORF on cosmid ZK154)
mec-7 Gene name (the 7 th “mechanosensory abnormality” gene named)
mec-7(e1506) Allele name (from the MRC Laboratory of Molecular Biology - e )
MEC-7 Protein name (product of mec-7 gene)
Mec Phenotype (Mechanosensory abnormal phenotype)
e1506 Homozygous allele
e1506/+ Heterozygous allele
mnDp30 Duplication (from the Herman Lab - mn )
nDf6 Deficiency/Deletion (from the Horvitz Lab - n )
muIs35 Integrated transgene (from the Kenyon Lab - mu )
evEx1 Extrachromosomal transgene array (from the Culotti Lab - ev )
CB3270 Strain name (from MRC Laboratory of Molecular Biology - CB)
mec-7p::gfp GFP transcriptional fusion (using only the promoter of the gene)
mec-7::gfp GFP translational fusion (in which gfp is inserted in the coding sequence of the gene)
ceh-6(pk33::Tc1) Transposon (Tc1) insertion in ceh-6 gene
1 All C. elegans gene names, allele designations, and reporter genes are written in italics. Cosmids, proteins, phenotypes, and strain names are not written in italics.
Mutation (allele) names : The wild-type allele of any gene is designated by an italicized plus sign, + . Mutant alleles are represented by 1-3 lowercase letters, which indicate specific laboratories, followed by a number. All gene and allele symbols are italicized, e.g., unc-54 , e678 , and mn5 . The homozygous genotype is represented by a single copy of the allele name ( e678 ). A heterozygous condition is indicated with a slash, e.g., e364/+ , e364/e1099. Chromosomal abnormalities are indicated by one or two letters after the lab code, e.g., mnDp30 is a duplication, nDf6 is a deletion (called a deficiency), and szT1 is a translocation. One or two letters may be added after the allele name to indicate particular properties conveyed by the mutation e.g. temperature-sensitivity (ts), amber (am), revertant (r), dominant (d), and semidominant (sd). When a gene has more than one mutation, such as could result from the intragenic reversion of a mutant allele, both the old and new mutations are indicated, e.g., intragenic reversion of e1498 could yield e1498u124r .
Gene names : Genes are designated by 3 or 4 lower case letters, a dash, and a number (all italicized), e.g. lon-2 and ensh-1 . To distinguish alleles of the same gene, the allele names are placed in parenthesis with no space between the gene and allele name, e.g., mec-7(e1343) and mec-7(e1506) . Genes that have been identified as an open reading frame (ORF) through bioinformatics approaches get a systematic gene identification ( e.g. ZK154.3) until subsequent studies prompts them to be given a gene name. The upstream promoter region of a gene is indicated by the gene name followed by a “p.” The promoter and the protein-coding names are separated by two colons as are parts of fusion proteins. Thus, a gene encoding a MEC-7::GFP translational fusion driven from the mec-7 promoter would be mec-7::gfp , whereas a mab-5 transcriptional fusion would be mab-5p::gfp . A transposon insertion into a gene is similarly shown using the transposon name and two colons, e.g., unc-22::Tc1 [most C. elegans transposons are labeled Tc (transposon, Caenorhabditis ) and a number].
Phenotypes : Mutant phenotypes are designated by the non-italicized gene name without the dash and number and with the first letter capitalized, e.g., Unc or Sma animals are Uncoordinated or Small in body size.
Gene products : RNAs are designated by the italicized gene name proteins are designated by the gene name in all caps and not italicized (MEC-7). Sometimes specific amino acid changes are indicated.
Strain names : Two capital letters and a number, e.g. CB429 and TU38, designate a strain containing one or more genetic differences. The letters indicate the laboratory that constructed the strain (http://www.cbs.umn.edu/research/resources/cgc/lab-head). None of these symbols are italicized. Because a given strain can carry more than one mutation, strain names can be thought of as shorthand to describe animals carrying complicated genotypes. Genes (and/or alleles) in strains with several mutations are listed in chromosome and then map order with genes on the same chromosome separated by spaces and genes on different chromosomes separated by semi-colons. The italicized name of the chromosome can also be included, e.g., lon-2(e678) mec-7(e1506) X , unc-54 myo-3 , and e364 e66 u38/szT1 . szT1 is one of many balancer chromosomes that are often used to maintain mutations that are lethal as homozygotes but viable as heterozygotes. For strains that contain reporter genes, there are two typical designations. The reporters are named for specific laboratories similar to mutant alleles, and they are further named according to whether the reporter gene is maintained extrachromosomally on a multicopy array using an “Ex” label ( e.g. evEx1) or is maintained by integrating the sequence (in most cases randomly) into the genome using an “Is” label ( e.g. muIs35 ).

Males are important because they allow the exchange of genetic material needed to generate animals with different genetic compositions and to map genes. Indeed, the animal has evolved to take advantage of the genetic contribution of rare males by using male (outcross) sperm before using hermaphrodite (self) sperm (Ward and Carrel 1979). Thus, if males are capable of mating, cross progeny prevail (to help males find hermaphrodites, researchers use Petri plates with a small spot of bacteria).

2.3. Life cycle

C. elegans embryogenesis takes approximately 16 hours at 20° (all of the subsequent times are also for development at 20°) (Figure 2). A virtually impermeable eggshell is made after fertilization, allowing the embryo to develop completely independent of the mother. However, embryos are usually retained within the hermaphrodite until about the 24-cell stage at which time they are laid. The hermaphrodite embryo hatches with 558 nuclei (some nuclei are in multi-nuclear syncytia, so the cell count is lower) and becomes a first stage (L1) larva. The animals begin to eat and develop through four larval stages (L1-L4). The L1 stage is

16 hr long the other stages are

12 hr long. Each stage ends with a sleep-like period of inactivity called lethargus (Raizen et al. 2008) in which a new cuticle (outer collagenous layer) is made. Lethargus ends with the molting of the old cuticle. Approximately 12 hr after the L4 molt, adult hermaphrodites begin producing progeny for a period of 2-3 days until they have utilized all of their self-produced sperm additional progeny can be generated if the sperm-depleted hermaphrodite mates with a male. After the reproductive period, hermaphrodites can live several more weeks before dying of senescence.

When bacteria are depleted and the animals are crowded, L2 larvae activate an alternative life cycle (Hu 2007) and molt into an alternative L3 larval stage called the “dauer” larva (“dauer” in German means “lasting" the signal is actually processed by L1 animals, but its results are not seen until the so-called “L2d” stage Golden and Riddle 1984). The dauer larva cuticle completely surrounds the animal and plugs the mouth preventing the animal from eating and thereby arresting development. The dauer cuticle has enhanced resistance to chemicals, so it provides the dauer with greater protection against environmental stresses and caustic agents. Dauer larvae can survive for many months and are the dispersal form most commonly encountered in the wild. When the dauer larvae are transferred onto plates with bacteria, they shed their mouth plugs, molt, and continue their development as slightly different L4 larvae.


Expansion microscopy of C. elegans

4.5x linear expansion) by isotropic swelling of chemically processed, hydrogel-embedded tissue. ExM of C. elegans is challenged by its cuticle, which is stiff and impermeable to antibodies. Here we present a strategy, expansion of C. elegans (ExCel), to expand fixed, intact C. elegans. ExCel enables simultaneous readout of fluorescent proteins, RNA, DNA location, and anatomical structures at resolutions of

65-75 nm (3.3-3.8x linear expansion). We also developed epitope-preserving ExCel, which enables imaging of endogenous proteins stained by antibodies, and iterative ExCel, which enables imaging of fluorescent proteins after 20x linear expansion. We demonstrate the utility of the ExCel toolbox for mapping synaptic proteins, for identifying previously unreported proteins at cell junctions, and for gene expression analysis in multiple individual neurons of the same animal.

Keywords: C. elegans developmental biology expansion microscopy gene expression immunohistochemistry in situ hybridization neuroscience super-resolution imaging synapses.

Conflict of interest statement

CY, AW, AS, FC Is a co-inventor on patents related to expansion microscopy: WO2015127183A2, US20170089811A1, US20160305856A1, US20170067096A1, US20160304952A1, WO2017147435A1, US20190256633A1, and WO2020013833A1, NB, AB, SA, CZ, MG, GH No competing interests declared, OH Reviewing editor, eLife, EB Is a co-inventor on patents related to expansion microscopy: WO2015127183A2, US20170089811A1, US20160305856A1, US20170067096A1, US20160304952A1, WO2017147435A1, US20190256633A1, and WO2020013833A1 and a co-founder of a company seeking to commercialize medical applications of expansion microscopy.

Figures

Figure 1.. Workflow for expansion of C.…

Figure 1.. Workflow for expansion of C. elegans (ExCel) sample processing.

Figure 2.. ExCel enables antibody-mediated visualization of…

Figure 2.. ExCel enables antibody-mediated visualization of fluorescent proteins.

( A ) Representative images of…

Figure 3.. Isotropy of ExCel.

Figure 3.. Isotropy of ExCel.

( A ) Representative images of paraformaldehyde-fixed, β-mercaptoethanol-reduced, AcX-treated, and…

Figure 3—figure supplement 1.. Local distortion at…

Figure 3—figure supplement 1.. Local distortion at the gonad region of day 1 – day…

Figure 3—figure supplement 2.. Local distortion at…

Figure 3—figure supplement 2.. Local distortion at the mouth region.

Representative images of mouth regions…

Figure 4.. Post-ExCel NHS-ester staining reveals anatomical…

Figure 4.. Post-ExCel NHS-ester staining reveals anatomical structures.

Representative images of ( A ) pharyngeal…

Figure 5.. ExCel enables simultaneous readout of…

Figure 5.. ExCel enables simultaneous readout of fluorescent proteins, RNA, DNA location, and anatomical features.

Figure 6.. Super-resolution imaging of synaptic proteins…

Figure 6.. Super-resolution imaging of synaptic proteins with ExCel.

( A ) Representative images of…

Figure 7.. Super-resolution imaging of electrical synapses…

Figure 7.. Super-resolution imaging of electrical synapses with ExCel.

Representative images of TagRFP-fused innexin protein…

Figure 8.. RNA detection in neurons.

Figure 8.. RNA detection in neurons.

ExCel-processed (formaldehyde-fixed, β-mercaptoethanol-reduced, LabelX- and AcX-treated, hydrogel-embedded, Proteinase-K digested…

Figure 9.. RNA detection in neurons, at…

Figure 9.. RNA detection in neurons, at sub-cellular resolution.

Representative images of ExCel-processed (formaldehyde-fixed, β-mercaptoethanol-reduced,…

Figure 10.. Single-neuron resolution RNA quantification.

Figure 10.. Single-neuron resolution RNA quantification.

( A ) A representative ExCel-processed (formaldehyde-fixed, β-mercaptoethanol-reduced, LabelX-…

Figure 11.. Screen of post-gelation treatments that…

Figure 11.. Screen of post-gelation treatments that confer tissue expandability and general stainability of epitopes.

Figure 12.. Immunohistochemistry after selected post-gelation treatments.

Figure 12.. Immunohistochemistry after selected post-gelation treatments.

( A ) Estimated signal-to-noise ratio of immunostaining…

Figure 13.. Workflow for epitope-preserving expansion of…

Figure 13.. Workflow for epitope-preserving expansion of C. elegans (epitope-preserving ExCel) sample processing.

70% estimated from the immunostaining results from the panel of 12 non-IgM antibodies in Figure 12A). Sample processing prior to Panel A is identical to the workflow for the standard ExCel protocol without ExFISH (as shown in blue arrows in Figure 1) until, and including, the gelation step (Figure 1A–C, E–G). The linear expansion factor of the hydrogel-specimen composite is shown in parentheses. For stages in which the worm tissue expands to a less extent than the surrounding hydrogel, which occurs due to incomplete homogenization of mechanical strength of the fixed worm tissue, the expansion factors of the worm and of the hydrogel are shown in front of and after a slash sign, respectively. (A–L) Steps of the protocol, with the bold text indicating the title of the step. (A) Hydrogel polymerization is performed on the specimen, by first incubating the specimens in activated monomer solution (0.015% 4-hydroxy-TEMPO, 0.2% TEMED, 0.2% APS, in addition to the non-activated monomer solution) for 1 hr at 4°C, transferring the specimens into a gelation chamber, and incubating the chamber for 2 hr at 37°C. During polymerization, AcX-modified proteins are covalently anchored to the hydrogel network. (B) Specimens are treated with chromatography-purified collagenase type VII at 0.5 kU/mL, in a calcium-containing tris-buffered saline (100 mM Tris pH 8.0, 500 mM NaCl, 40 mM CaCl2) overnight (18–24 hr) at 37°C. During this treatment, the hydrogel expands by

1.2x linearly, whereas the worm slightly reduces in size to

0.9x linearly. Due to the mismatch in expansion factor between the worm and the gel, the worm tissue detaches from the surrounding hydrogel, but physically remains in the hydrogel mold that was made of its own shape during the gelation step in A. (C) Specimens are processed with a denaturation treatment, in which they are incubated in a minimally-expanding protein-denaturing buffer (MAP5 buffer 50 mM Tris pH 9.0, 5.72% (w/w) sodium dodecyl sulfate (SDS), 400 mM NaCl, 20 mM CaCl2) overnight (18–24 hr) at 37°C, overnight (18–24 hr) at 70°C, and 2 hr at 95°C. Reduced calcium and NaCl concentrations are used in this buffer, compared to other non-expanding buffers designed in this paper, due to their incompatible solubility with SDS at higher concentrations. (D) Specimens are washed four times in a tris-buffered saline (TNC40020 (acronyms are used in the supplementary protocols in Appendix 1) buffer 50 mM Tris pH 8.0, 400 mM NaCl, 20 mM CaCl2) to remove SDS from the hydrogel sample. Specimens are then washed four times in tris buffered saline with reducing calcium concentration (once with TNT Buffer + 10 mM CaCl2, and then three times with TNT Buffer TNT Buffer is 50 mM Tris pH 8.0, 1M NaCl, 0.1% Triton X-100) to remove calcium ions from the hydrogel sample. Finally, specimens are washed with phosphate-buffer saline with reducing NaCl concentration (once with PBST + 500 mM NaCl, twice with PBST PBST is 1x PBS + 0.1% Triton X-100). (E) Specimens are immunostained with fluorescent antibodies against the target antigens. (F) Specimens are incubated with AcX at a concentration of 0.1 mg/mL in PBST (1x PBS + 0.1% Triton X-100) overnight at RT. This step equips proteins, including the fluorescent antibodies introduced in E, with a polymer-anchorable moiety. (G) Specimens are incubated in non-activated G2 monomer solution (50 mM MOPS pH 7.0, 2 M NaCl, 7.5% (w/w) sodium acrylate, 2.5% (w/w) acrylamide, 0.15% (w/w) N,N’-methylene-bis-acrylamide) overnight at 4°C, to ensure complete diffusion of the monomer solution throughout the specimen, prior to the gelation reaction. (H) Specimens are re-embedded into a second expandable hydrogel, by incubating the specimens in activated monomer solution (0.015% 4-hydroxy-TEMPO, 0.2% TEMED, 0.2% APS, in addition to the non-activated monomer solution) for 45 min at 4°C, transferring the specimens into a gelation chamber, and incubating the chamber for 2 hr at 37°C. During polymerization, AcX-modified fluorescent antibodies are covalently anchored to the hydrogel network of the second hydrogel (orange grids). We use blue grids to represent the hydrogel network of the first, DATD-crosslinked hydrogel (i.e., the network synthesized in Panel A), to differentiate it from the network of the re-embedding second hydrogel. (I) Specimens are treated with Proteinase K at 8 U/mL, in non-expanding digestion buffer (50 mM Tris pH 8.0, 500 mM NaCl, 40 mM CaCl2, 0.1% Triton X-100) overnight (18–24 hr) at RT, to further reduce the mechanical strength of the original worm tissue and permit greater expansion. During this proteolytic treatment, most proteins lose antigenicity, but some of the fluorescent signals from AcX-anchored fluorescent proteins are retained, as utilized by the original ProExM protocol. (J) Specimens are treated with DATD cleaving solution (20 mM sodium meta-periodate in 1x PBS, pH 5.5) for 30 min at RT, to chemically disintegrate the first hydrogel, which contains the periodate-cleavable crosslinker N,N'-diallyl-tartardiamide (DATD), while sparing the second hydrogel, which contains a periodate-resistant crosslinker, N,N’-methylene-bis-acrylamide (bis). (K) To visualize anatomical features, specimens can be stained with an N-hydroxysuccinimide ester (NHS ester) of fluorescent dye (introduced in Main Text, Figure 4, Videos 1 and 2). NHS-ester staining is performed at 5 μM in PBST (1x PBS + 0.1% Triton X-100) overnight at RT. DAPI staining can be applied at this stage, but the result does not correspond to the expected nuclear pattern as observed in Figures 2, 5 and 9 (see Figure 13—figure supplement 1). (L) Specimens are expanded with one wash in 0.1x PBS and two washes in deionized water. At this stage, the hydrogel expands by

7.4x linearly, whereas the worm tissue expands by

2.8x linearly, within a range from 2.5 to 3.5x (median, 2.78x mean, 2.83x n = 10 independently processed hydrogels from 2 sets of experiments). After expansion, specimens are ready for imaging.

Figure 13—figure supplement 1.. DAPI staining after…

Figure 13—figure supplement 1.. DAPI staining after epitope-preserving ExCel.

Representative images of epitope-preserving-ExCel-processed (formaldehyde-fixed, β-mercaptoethanol-reduced,…

5 μm, in biological units) of the tissue, and not across the entire animal, in order to facilitate visualization of the pattern of staining (since the max projection across the entire animal would contain signals from overlapping layers of nuclei, which hinders interpretation). Brightness and contrast settings: first set by the automatic adjustment function in Fiji, and then manually adjusted (raising the minimum-intensity threshold and lowering the maximum-intensity threshold) to improve contrast. Linear expansion factor: worm, 3.0–3.2x surrounding hydrogel, 7.6x. Scale bars: (A, C) 5 μm (B, D) 1 μm (in biological units, i.e. post-expansion lengths are divided by the expansion factor of the worm).

Figure 14.. Isotropy of epitope-preserving ExCel.

Figure 14.. Isotropy of epitope-preserving ExCel.

( A ) Representative images of paraformaldehyde-fixed, β-mercaptoethanol-reduced, AcX-treated,…

Figure 15.. Epitope-preserving ExCel allows multiplexed imaging…

Figure 15.. Epitope-preserving ExCel allows multiplexed imaging of endogenous proteins at nanoscale resolution.

Figure 16.. Super-resolution imaging of pre-synaptic active…

Figure 16.. Super-resolution imaging of pre-synaptic active and peri-active zones with epitope-preserving ExCel.

Figure 17.. Workflow for iterative expansion of…

Figure 17.. Workflow for iterative expansion of C. elegans (iExCel) sample processing.

20x. Sample processing prior to Panel A is identical to the workflow for the standard ExCel protocol without ExFISH (as shown in blue arrows in Figure 1) until, and including, the post-Proteinase-K partial expansion step (Figure 1A–C, E–G). The linear expansion factor of the hydrogel-specimen composite is shown in parentheses. (A–I) Steps of the protocol, with the bold text indicating the title of the step. (A) Specimens are partially expanded from a linear expansion factor of 1.0x to 1.8x, with the same protocol as shown in Figure 1I. (B) Specimens are immunostained first with primary antibodies against fluorescent proteins in 5x SSCT overnight at 4°C, and then with secondary antibodies that have been conjugated to a 24-base DNA oligonucleotide, in DNA-conjugated Antibody Staining Buffer (2x SSC, 2% (w/v) dextran sulfate, 1 mg/mL yeast tRNA, 5%(v/v) normal donkey serum, 0.1% Triton X-100) overnight at 4°C. The DNA oligo is conjugated to the antibody at the 3’ end, and contains a gel anchorable group at the 5’ end. (C) Specimens are expanded from a linear expansion factor of 1.8x to 3.8x, with the same protocol as shown in Figure 1M. (D) Specimens are re-embedded into another non-expandable hydrogel (‘Gel #2’) to lock up its size at the expanded state, as shown in Figure 1N, except that the monomer solution is replaced by DATD-crosslinked re-embedding monomer solution (10% acrylamide, 1% N,N'-diallyl-tartardiamide (DATD), 0.05% TEMED, 0.05% APS), which results in a hydrogel that can be later disintegrated via crosslinker cleavage, to allow full expansion of the final expandable gel. The DATD-crosslinked re-embedding monomer solution contains a charged molecule APS. Therefore, the linear expansion factor slightly drops from 3.8x to 3.6x during this step. During hydrogel polymerization, the DNA oligo on the antibody, which contains a gel-anchorable group, is covalently anchored to the second hydrogel network (orange grids). (E) Specimens are incubated with a 100-base DNA oligonucleotide (‘Linker’), which hybridizes to the 24-base DNA oligo on the secondary antibodies, and which contains a gel anchorable group on its 5’ end, in iExCel hybridization buffer (4x SSC, 20% (v/v) formamide) overnight at RT. (F) Specimens are re-embedded into another expandable hydrogel (‘Gel #3’), by incubating the specimens in activated Gel #3 monomer solution (1x PBS pH 7.4, 7.5% (w/w) sodium acrylate, 2.5% (w/w) acrylamide, 0.15% (w/w) N,N’-methylene-bis-acrylamide, 2M NaCl, 0.015% 4-hydroxy-TEMPO, 0.2% TEMED, 0.2% APS) for 50 min at 4°C, transferring the specimens into a gelation chamber, and incubating the chamber for 2 hr at 37°C. During polymerization, the linker DNA oligo, which contains a gel-anchorable group, is covalently anchored to the hydrogel network of the third hydrogel (magenta grids). (G) Specimens are treated with DATD cleaving solution (20 mM sodium meta-periodate in 1x PBS, pH 5.5) for 30 min at RT, to chemically disintegrate the first and the second hydrogels, which contain a periodate-cleavable crosslinker N,N'-diallyl-tartardiamide (DATD), while sparing the third hydrogel, which contains a periodate-resistant crosslinker N,N’-methylene-bis-acrylamide (bis). (H) Specimens are incubated with a fluorophore-conjugated 15-base locked nucleic acid (LNA) oligonucleotide, which hybridizes to the 100-base linker DNA oligo at four locations, in iExCel hybridization buffer (4x SSC, 20% (v/v) formamide) overnight at RT. (I) Specimens are expanded to a linear expansion factor of

20x, with three washes in deionized water. After expansion, specimens are ready for imaging.


Introduction

Caenorhabditis elegans is an important model system in biology, because of its tractable size (959 somatic cells in adult hermaphrodites), its genetic manipulability, and its optical transparency, which yields the possibility of whole-organism imaging of biological processes and signals. Perhaps not surprisingly, therefore, super-resolution microscopy has been useful to the analysis of C. elegans, with studies applying STORM, PALM, SR-SIM, and STED to C. elegans to investigate cells and tissues in both intact or dissected C. elegans (Rankin et al., 2011 Gao et al., 2012 Vangindertael et al., 2015 He et al., 2016 Köhler et al., 2017 Krieg et al., 2017). However, the depths of imaging of such studies were largely physically limited to a few microns to tens of microns, insufficient to map the entire depth of an adult animal, and the hardware required for super-resolution microscopy is not available in all laboratories, and can be slow and/or expensive to deploy. Furthermore, the tough cuticle of C. elegans presents a barrier to immunostaining in the intact animal, important for STORM and STED imaging and for the general labeling of proteins in a variety of scientific contexts.

Recently, we discovered that it is possible to isotropically expand biological specimens by permeating them evenly and densely with a swellable hydrogel polymer network, anchoring key biomolecules or labels to the hydrogel, softening the tissue through a chemical process, and then adding water, which swells the polymer and in turn the tissue (Chen et al., 2015). This technique, expansion microscopy (ExM), is now being adapted and improved by many groups, and has been applied to tissues of mice, human patients, and in many other biological contexts (Chen et al., 2015 Chen et al., 2016 Chozinski et al., 2016 Ku et al., 2016 Tillberg et al., 2016 Chang et al., 2017 Zhao et al., 2017 Park, 2018 Truckenbrodt et al., 2018 Gambarotto et al., 2019 Wassie et al., 2019). However, C. elegans is wrapped in a multi-layer cuticle, which is well known to be impermeable to many small molecules and all antibodies, and mechanically stiff to the point where physical expansion would be expected to proceed poorly (Duerr, 2006 Page and Johnstone, 2007 Chisholm and Xu, 2012). Thus, we set out to develop an ExM protocol customized for the C. elegans context that would overcome these barriers.

To achieve this goal, we modified previously published protocols in a number of ways (Figure 1, green steps) to generate a new protocol which we call expansion of C. elegans (or ExCel). This protocol results in high signal-to-background antibody staining against protease-resistant fluorescent proteins, low-distortion (

1–6% over length scales of 0–100 μm) physical expansion by

3.3x, and both protein and RNA detection with sub-cellular resolution. Using ExCel, we were able to resolve synaptic and gap junction proteins better than with ordinary confocal microscopy, and simultaneously image proteins, RNA, and DNA location within the same specimen. In particular, such multiplexed capability has not been demonstrated with previous super-resolution methods in C. elegans, and facilitates nanoscale-precise analyses of how multiple molecular types are spatially organized in the context of an entire animal.

Workflow for expansion of C. elegans (ExCel) sample processing.

A method for expanding cuticle-enclosed intact C. elegans, extending published proExM and ExFISH protocols with specific modifications (shown in green text full key in lower left). Depending on whether the user intends to visualize RNAs or not, the protocol branches into two forms. The protocol without ExFISH, which supports the readout of fluorescent proteins, DNA location (in the form of DAPI staining), and anatomical features, is indicated with blue arrows, ending in Panel L. The protocol with ExFISH, which additionally supports readout of RNAs, is indicated with orange arrows, ending in panel Q. For all steps after hydrogel formation (Panels G-Q), the linear expansion factor of the hydrogel-specimen composite is shown in parentheses. (A–Q) Steps of the protocol, with the bold text indicating the title of the step see text for details of each step.

The standard ExCel protocol visualizes fluorescent reporters, such as those fused to proteins of interest, which requires transgenesis, and could in principle affect the function and localization of the target protein. Thus, we additionally developed an alternative ExCel protocol, which we call epitope-preserving ExCel, that enables detection of untagged, completely endogenous proteins, using off-the-shelf primary antibodies. The epitope-preserving ExCel protocol replaces the use of Proteinase K, a general protease that disrupts most epitopes in the standard ExCel protocol, with an epitope-preserving cuticle-permeabilization treatment that we identified in a systematic screen of chemical treatments. This protocol enables antibody staining of protein epitopes at the expense of a slightly reduced expansion factor (

2.8x) and lower expansion isotropy (

8–25% error over length scales of 0–100 μm). We showed that epitope-preserving ExCel allows multiplexed readout of multiple native proteins at super-resolution, a capability that we used to identify a previously unreported protein localization at the junctions between developing vulval precursor cells, and to resolve the peri-active and active zones of chemical pre-synapses.

Lastly, we developed a third protocol, iterative ExCel (iExCel), which enables two successive rounds of hydrogel-mediated expansion of a given worm, by incorporating the previously validated strategies of iterative expansion microscopy into the ExCel context (Chang et al., 2017). iExCel brings the expansion factor from

20x, and the theoretical limit of resolution down to

25 nm, at a low level of distortion (

1.5–4.5% over length scales of 0–100 μm), on par with that of standard ExCel, on which it builds. With iExCel, we were able to resolve fluorescent puncta that may represent individual GFP molecules expressed in the neuronal cytosol.

Each of these ExCel protocols highlights some of the challenges remaining in deploying ExM in C. elegans, including distortion in the gonad and mouth regions, reduced general isotropy with epitope-preserving ExCel, and the ability to only detect fluorescent proteins with the current form of iExCel, which provide grounds for further optimization in the future.


Talk Overview

Can a single human brain ever understand how a human brain works? Colón-Ramos argues not – but a network of brains might! Networks are an essential part of science, whether it is at the level of single neurons connecting to form a brain, or networked brains forming a scientific community. Colón-Ramos studies neural networks in the nematode C. elegans to uncover how neurons organize and communicate with each other to form the nervous system. Using the nervous system as a metaphor, he explains how our own brains, similar to neurons, are “wired to be wired”, and how scientists connect their brains to weave new knowledge.


Supporting information

S1 Fig. csGPCR gene locus analysis.

(A) Histogram of upstream intergenic region distances of all C. elegans csGPCR genes.

The average size of the 5’ intergenic region (= distance to next gene) is 1.8 kb.

Eighty-nine percent of all loci have a 5’ intergenic region smaller than 4 kb.

(B) Histogram of average combined intron length (bp) per GPCR gene.

(C) The intergenic region of the majority of GPCR is substantially larger

than the combined intronic region. bp, base pair csGPCR, chemosensory-type GPCRs GPCR, G-protein-coupled receptor.

S1 Table. Masterlist of all examined GPCR reporters.

GPCR, G-protein-coupled receptor protein.

S2 Table. List of all identified sensory neurons with GPCR expression.

Gene in bold: newly identified in this paper

Gene in non-bold: previously identified

(Gene in parenthesis): ID based on position and morphology, not confirmed with neuron-specific reporter.

GPCR, G-protein-coupled receptor protein.

S3 Table. Primers.

Primer sequences for the reporters generated by the Vancouver consortium (BC strains) can be found at http://www.gfpworm.org.


H1N1? H2N5? What Do Flu Names Mean?

The fall brings with it cooler days, apple picking and flu season.

We know what the symptoms of the flu are, how it is spread and the importance of getting a flu shot. What we may not know is the difference between the strains of the influenza virus and why are they referred to by the letters H and N - followed by numbers.

This may be a bit "inside biology" for some, but, in case you are wondering what these names mean and how they come about, here is a look into the virology behind the influenza virus.

Strains of influenza are characterized by two proteins that are on the outer surface of the virus: hemagglutinin and neuraminidase. This is where the "H" and "N" in the name come from. The proteins can be seen in the picture above, represented by the blue "spikes" on the outside of the virus. Both of these proteins are required for the virus to cause an infection and perform complementary functions. The hemagglutinin is critical for the virus to be able to attach to, and then enter the cell. Without hemagglutinin, the entire process of infection could not be initiated.

Once in the cell, the virus takes over the normal cell machinery and uses it to make many copies of itself.

Then, the neuraminidase enzyme comes into play. Neuraminidase is required for the newly made viruses to escape the host cell, where they can then perpetuate the infection. It's role is to clip the newly made viruses from the membrane of host cell and release them. Without neuraminidase, the new viruses would stay attached to the host cell, unable to infect new cells.

When a person becomes infected with influenza virus, their body's immune system responds by making antibodies to the H and N proteins. Then the antibodies bind to the H and N proteins, and block them from doing their job, stopping the virus in its tracks.

However, with influenza virus, the situation is even more complicated.

Influenza's complexity starts with the fact that there are 14 versions of H protein and 9 versions of N which means that there are a total of 144 varieties of flu. Not all of these 144 are infective.

Additionally, there are two major processes that create a complicated situation even worse. These are called antigenic drift and antigenic shift.

Antigenic drift means that either H or N change in a particular strain. This results in new strains, and this trips up the immune system. When H and N mutate, it is possible that they change so much that the antibodies may not bind to them anymore, leaving the virus fully infective.

The concept of antigenic shift is more complicated, but, stay with me for a moment. In order to understand antigenic shift, first we must understand how the genome of influenza is stored inside the virus. Influenza belongs to the orthomyxoviridae family of viruses. This means that, unlike our genes, which are made up of DNA, the flu virus's genome is made up of RNA - eight separate pieces of RNA. Flu is an example of an RNA virus with the hemagglutinin and neuraminidase genes are found on different pieces of RNA.

When two different strains of flu infect a cell at the same time, the pieces of RNA (eight from each virus) mix together. This is called reassortment and results in a new strain.

In looking at the figure below, let's say that strain A (with the green RNA) is an H1N1 virus and strain B (with the blue RNA) is an H2N5. In the figure, both of these viruses are infecting the same cell at the same time. When in the cell, the 16 pieces of RNA mix together and the virus that is released from the cell is different (a combination of some green RNA and some blue RNA) from either of the original ones that went in.

This process makes pandemics more likely because the strain that results from the reassortment may be a more dangerous strain and also one that no one has been exposed to before.

Within the past century, there have been four flu pandemics (worldwide epidemics.) They occurred in 1918 (H1N1 - swine flu), 1957 (H2N2), 1968 (H3N2) and 2009 (H1N1) and experts agree that it is not a matter of if, but when, the next pandemic will strike. It is not simple to predict which flu strain will be wreaking havoc next and the flu vaccine varies in how effective it is from year to year. Knowing about the complexity of the influenza virus's genome and its ability to undergo antigenic shift and drift help us understand, at least to some extent, what we are up against each year.


AUTHOR CONTRIBUTIONS

Erick O. Olivares: Data curation Formal analysis Methodology Software Visualization Writing – original draft Writing – review & editing. Eduardo J. Izquierdo: Conceptualization Formal analysis Funding acquisition Investigation Methodology Project administration Supervision Visualization Writing – original draft Writing – review & editing. Randall D. Beer: Conceptualization Formal analysis Funding acquisition Investigation Methodology Project administration Resources Software Supervision Writing – original draft Writing – review & editing.


Watch the video: Vulval Development in (January 2022).