Collection and maintenance of A. suum worms

Adult female A. suum worms were collected from the JBS Swift and Co. pork processing plant at Marshalltown, IA, and were maintained in Ascaris Ringers Solution (ARS: 13 mM NaCl, 9 mM CaCl₂, 7 mM MgCl₂, 12 mM C₄H₁₁NO₃/ Tris, 99 mM Na acetate, 19 mM KCl and 5 mM glucose pH 7.8) at 32°C for 24 hrs. to allow for acclimatization before use in experiments. The solution was changed twice daily, and worms were used within three days of collection for experiments. All worms were examined at the start of each day and were excluded if they were damaged or became immotile.

Injection of compounds into Ascaris suum

Healthy and motile A. suum were selected and placed in a dissection tray filled with warmed ARS. Worms were left to acclimatize in the tray for two mins and were monitored for movement. All solutions were prepared in Ascaris Perienteric Fluid APF (23 mM NaCl, 110 mM Na acetate, 24 mM KCl, 6 mM CaCl₂, 5 mM MgCl₂, 5 mM HEPES, 11 mM D-glucose, pH 7.6 using NaOH or acetic acid). Worms were injected with either 500 µL APF (negative control), 500 µL 30 µM levamisole (positive control), 500 µL 30 µM betaine or 500 µL 30 µm β-alanine betaine. Solutions were injected into the worm above the collar using a 23-gauge needle attached to a 1 mL Luer slip syringe. Worms were monitored for 2 mins and were assessed for paralysis.

A. suum cDNA synthesis and RT-PCR detection of DEG-3/DES-2 nAChR superfamily members and Sanger sequencing

A. suum orthologues of the DEG-3/DES-2 family members and LGC-41 were identified by blasting the C. elegans protein and cDNA sequences for each target, obtained from WormBase (www.wormbase.org ver. WS298), against the A. suum genome (PRJNA62057) found on WormBase Parasite (parasite.wormbase.org ver. WS291). The identified orthologues for each gene and the accession number for ENA depository are presented in Table 2.

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Table 2. Genes and accession numbers for members of DEG-3/DES-2 nAChR superfamily in A. suum. Note Asu-deg-3 and Asu-des-2 had strong similarity with AgR001_g107 but had different predicted isoforms.

https://doi.org/10.1371/journal.pntd.0014518.t002

The A. suum intestine was separated from the body wall, which includes muscles, muscle bags and nerves, and both tissues were homogenized separately in 1 mL of Trizol reagent using a mortar and pestle, followed by total RNA extraction according to the Trizol Reagent protocol (Life Technologies, Waltham, MA, USA). One microgram of total RNA from each tissue was used to generate cDNA by reverse transcription (RT) using SuperScript VILO Master Mix (Life Technologies, Waltham, MA, USA) following the manufacturer’s protocol. RT-PCR was conducted to detect the presence of Asu-acr-20, Asu-acr-23, Asu-deg-3, Asu-des-2, and Asu-lgc-41 using primers targeting coding regions of each gene (Table 1). Asu-gapdh was used as a reference gene. Negative controls included enzyme, water, and both forward and reverse primers for the target with no cDNA template. The cycling conditions for PCR were an initial denaturation for 2 min at 95 °C, followed by 35 cycles of 95 °C for 30 sec, 60 °C for 35 sec, 72 °C for 45 sec, and a final extension at 72 °C for 10 min using GoTaq G2 Hot Start Green Master Mix (Promega, USA). PCR products were separated on a 2% agarose gel containing SYBR Safe DNA Gel Stain (Invitrogen, Waltham, MA, USA) followed by visualization and images were captured using an Azure 600 imaging system (Azure Biosystems, Dublin, CA, USA) set to SYBR Safe (excitation 472 nm, emission 595 nm). Uncropped and unedited raw images are shown in S1 Raw gel.

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Table 1. Primer sequences used for RT-PCR. Forward and reverse primer sequences used for the detection of acr-20, acr-23, deg-3, des-2, lgc-41 and the reference gene gapdh in Ascaris suum. The primers used to amplify the full-length Asu-acr-23 construct are provided labelled FL.

https://doi.org/10.1371/journal.pntd.0014518.t001

Amplification of full-length Asu-acr-23 was performed using Platinum SuperFi II Green PCR Master Mix (Thermo Fisher Scientific, Waltham, MA, USA) under the following conditions: 98 °C 30s; 35 cycles at 98 °C 30 s, 60 °C for 20 s, 72 °C for 30 s and a final extension at 72 °C for 10 mins. Products were visually identified on a 1% agarose gel containing SYBR Safe as previously described. Target bands were excised and purified with the NucleoSpin Gel and PCR Clean-up kit (Macherey-Nagel, Düren, Germany) per the manufacturer’s instructions. Purified amplicons were submitted to the Iowa State University DNA Sequencing Facility for Sanger sequencing (bidirectional reads were achieved using the forward and reverse primers provided in Table 1). The full-length sequencing result is provided in S4 Fig.

C. elegans strains and maintenance

C. elegans strains were maintained on NGM agar plates seeded with E. coli OP50 bacteria per standard protocols [11]. Strains used were: N2, RB2119 acr-23(ok2804) V, VC1598 acr-20(ok1849)/mT1 II; + /mT1 [dpy-10(e128)] III, TU1803 deg-3(u662); des-2(u695) V and PS8729 lgc-41(sy1494) X. All strains were acquired from the Caenorhabditis Genetics Center (University of Minnesota, Minneapolis, MN, USA).

Ascaris suum muscle flaps for electrophysiology

1 cm muscle tissue flaps were prepared by dissecting the anterior part of the worm, 2–3 cm caudal to the head. A body muscle flap preparation was then pinned onto a Sylgard-lined double jacketed bath chamber maintained at 35 ˚C by inner circulation of warm water using a 6-liter circulating Isotemp3016H water bath (Fisher scientific, Waltham, MA, USA). The intestine was removed to expose the muscle cells [12]. The preparation was continuously perfused, unless otherwise stated, with APF. The incoming perfusate was pre-warmed to 35˚C with an SH 27B in-line heating system (Warner Instruments, Holliston, MA, USA) before it was perfused at 3.5-4 mL/min through a 20-gauge (1.5″ long) needle placed directly over the muscle bag. Stocks of the experimental compounds were dissolved in APF. Concentrations of 30 µM, 100 µM, 300 µM and 1 mM β-alanine betaine were applied for a period of 10 secs by means of a tube that rapidly replaced all the bathing solution covering the Ascaris muscle cells. Flanking applications of 10 µM ACh were applied at the start and end of each recording for 10 secs to determine viability.

A single micropipette was used to record the membrane potential and to examine the electrophysiological effects in the A. suum muscle bag region. Borosilicate capillary glass (Harvard Apparatus, Holliston, MA, USA, ID-0.86mm, OD- 1.5mm) micropipettes were pulled on a Flaming Brown Micropipette puller (Sutter Instrument Co., Novato, CA, USA) and filled with 3 M potassium acetate (resistance 20–30 MΩ). The recordings were obtained by impaling the bag region of A. suum muscle with both the micropipettes. All experiments were performed using an Axoclamp 2A amplifier, a 1320A Digidata interface and Clampex 9 software (Molecular Devices, San Jose, CA, USA).

C. elegans paralysis assays

Motility responses to β-alanine betaine were assayed on NGM plates that were prepared four hours before the experiment. 50 mM β-alanine betaine was added to molten agar (~55°C). Once solidified, fresh E. coli OP50 was added to the plates, which were left to air dry (~ 2 hrs.). 20–40 L4 animals were picked the night before the assay onto fresh OP50 seeded NGM plates and left to develop to young adults overnight at room temperature. Animals were transferred from the stock plate to an intermediate plate for one minute to allow animals to separate and be assessed for motility. Ten suitable animals were transferred to β-alanine betaine assay plates and motility was measured. Worms that either appeared damaged or had reduced movement or no movement were removed from the assay plate and replaced with viable C. elegans. Animals were allowed to free roam on the bacterial lawn and were assessed after 10, 20, 30, 45, 60, 90 and 120 mins. Animals were scored based on movement by lightly tapping the ‘nose’ of the worm with a hair. If the worm failed to respond to the hair, they were considered paralyzed.

Preparation and loading Fluo-3AM

A 2 cm section of the A. suum intestine was removed from the body piece using fine forceps and cut open. The intestinal flap was placed into a laminar flow recording chamber (RC-26) which was secured on top of a coverslip (24 x 50 mm) in a PH-1 heated platform and pinned using a slice anchor (26 x 1mm x 1.5mm grid) (Warner Instruments, Holliston, MA, USA). The sample was immersed in 1 mM CaCl₂ APF (23 mM NaCl, 110 mM Na acetate, 24 mM KCl, 1 mM CaCl₂, 5 mM MgCl₂, 5 mM HEPES, 11 mM D-glucose; pH 7.6). Fluo-3AM (Sigma-Aldrich, St. Louis, MO, USA) loading was achieved by incubating the tissue in Ca²⁺-free (<100 µM) APF containing 5 µM Fluo-3AM and 0.05% Pluronic F-127 (10% v/v in water; Invitrogen, Waltham, MA, USA) for 60 mins with the recording chamber connected to a Dual Automatic Temperature Controller (Warner Instruments, Holliston, MA, USA) maintained at 36–37°C. After incubation, the Fluo-3AM solution was discarded, and the sample was incubated in 1 mM CaCl₂ APF for an additional 20 mins at 36–37°C to promote Ca²⁺ loading. Fluo-3AM loading was confirmed under blue light using a blue/green filter box (EF-4 AT EGFP/FITC/CY2/ALEXA FLUOR Band Pass; excitation: 465–495 nm, emission: 525–545 nm, dichroic mirror) (Chroma Technology, Bellows Falls, VT, USA) and visualized under pseudo-color settings using MetaFluor 7.10.2 (Molecular Devices, San Jose, CA, USA). Any tissue that did not show Fluo-3 fluorescence was discarded.

Measurement of Ca²⁺ fluorescence

All recordings were performed on a Nikon Eclipse TE3000 microscope (Nikon, Tokyo, Japan) fitted with a 20X/0.45 Nikon PlanFluor objective, using a PhotoMetrics Retiga R1 Camera (PhotoMetrics, Inc., Huntington Beach, CA, USA). Light control was achieved using a Lambda 10–2 with a shutter controller. Fluorescence was achieved using a Lambda LS Xenon bulb lightbox (Sutter Instruments, Novato, CA, USA) which delivered light via a fiber optic cable to the microscope which passed through the blue/green filter box. Fluorescent light emission was controlled by using the shutter. Minimal illumination exposure was used to prevent photobleaching.

During recordings, tissues were continuously perfused with 1 mM CaCl₂ APF. Intestinal preparations were exposed to 1 mM β-alanine betaine for five mins. Application of 10 mM CaCl_2, which was used as a positive control to determine tissue viability, was applied 5 mins after the β-alanine betaine signal returned to baseline to detect delayed or latent responses to occur. Every experiment reported in this study had a 100% response to 10 mM CaCl₂. A change in fluorescence ≥5% was classified as a positive response to a compound as the mean spontaneous change in cells continuously perfused with APF buffer was ~ 2.5% [13]. Stocks and working concentrations of all compounds were made in 1 mM CaCl₂ APF buffer solution.

All solutions were delivered to the chamber under gravity feed through solenoid valves controlled using a VC-6 six-channel valve controller through an inline heater set at 37°C (Warner Instruments, Holliston, MA, USA), at a rate of 1.5mL/min. At the start of all experiments, intestinal preparations were left under blue light for a minimum of 3 mins to promote settling and equilibration of the fluorescent signal and to monitor spontaneous Ca²⁺ signals.

All Ca²⁺ signal recordings were acquired and analyzed using MetaFluor 7.10.2 with exposure settings at 250 ms with 2x binning. Ca²⁺ signals from each intestine were collected from 50 square 50 µm x 50 µm areas across the intestine covering a total area of 125,000 µM² that included 800–1000 individual enterocytes. The mean fluorescence amplitude was calculated for each intestinal exposure of all 50 regions. Maximal percent Ca²⁺ signal amplitudes (ΔF) were calculated using the equation F1-F0/F0 x 100, where F1 is the fluorescent value and F0 is the baseline value. All F0 values were taken at the time point each compound was applied for every region analyzed. All representative response traces are presented as the mean percent change in Ca²⁺ fluorescence with the standard error of the mean (±SEM) of all regions from a single recording. Traces were generated by converting the Ca²⁺ signal profiles of all the regions from a single recording to percentages using the previously described ΔF/F0 equation, with the time of stimulus application being F0 (0% for all regions) and 100% being the peak value for each region. The number of intestinal tissues recorded (n), along with the total number regional responses are provided in the figure legends.

Determination of choline analogues by HILIC–MS and HILIC–MS/MS

2.5 mL A. suum pseudocoelomic fluid samples were acquired from freshly collected worms and 20 µM neostigmine was added as an esterase inhibitor to reduce the hydrolysis of acetylcholine. Choline analogues in A. suum pseudocoelomic fluid were initially identified using hydrophilic interaction liquid chromatography coupled to full mass range (HILIC-MS). The presence of choline and β-alanine betaine were then confirmed using tandem mass spectrometry (HILIC–MS/MS).

Sample preparation

Fluid samples were maintained on ice and mixed thoroughly. Proteins were precipitated by adding three volumes of ice-cold acetonitrile. Samples were vortexed for 30 s and incubated on ice for 10 min. The mixture was centrifuged at 15,000 × g for 10 min at 4 °C. The supernatant was transferred to autosampler vials and diluted with acetonitrile to achieve 80–90% organic solvent prior to injection.

HILIC chromatography

Chromatographic separation was performed using a Thermo Fisher Scientific Vanquish Flex UHPLC system with a HILIC column (Agilent Infinity Lab Poroshell 120 HILIC-Z; 2.1 x 100 mm, 2.7 µm particle size) maintained at 30 °C. The mobile phases consisted of: A: 95:5 water/acetonitrile with 10 mM ammonium acetate and mobile phase and; B: 95:5 acetonitrile/water with 10 mM ammonium acetate. The gradient started at 90% B, ramped linearly to 50% B over 5 mins, then from 50% B to 40% B over 1 min, held for 2 mins at 40% B, then returned to initial conditions over 1 min, and equilibrated for 6 mins. The flow rate was 0.3 mL/min, and the injection volume was 10 µL.

Mass spectrometry

Detection was performed using a Thermo Fisher Scientific Exploris 120 quadrupole-orbitrap mass spectrometer equipped with an electrospray ionization (ESI) source operated in positive ion mode. The following source parameters were used: capillary voltage 3.5 kV, sheath gas = 50 (Arb), aux gas = 10 (Arb), sweep gas = 1 (Arb), ion transfer tube temp = 325 °C, vaporizer temp = 350 °C.

Additional mass spectrometer settings include: full mass range resolution = 60,000, MS/MS resolution = 15,000, full mass range = 50 – 500 m/z, MS/MS mass range was set to “auto”, Automatic Gain Control (AGC) was set to ‘standard’ for both full mass range and MS/MS, and maximum ion time was set to ‘auto’ for both full mass range and MS/MS. MS/MS data was acquired in t-MS2 (targeted MS2) mode.

Statistical analysis

Statistical analysis of all data was done using GraphPad Prism 9.0 (GraphPad Software, Inc., La Jolla, CA, USA). To ensure reproducibility, we repeated our experiments: the numbers of animals, number of preparations, the concentrations, and durations of applications are provided in the legends of the figures. Analysis of changes in depolarization was done using paired or unpaired student t-tests with P < 0.05 being considered significant. All data are presented as mean ± SEM for each treatment.

Chemicals

Source of chemicals: β-alanine betaine was produced by ISU and characterization matched previous reports [14]. For the mass spectrometry experiments, the β-alanine betaine neat chemical standard was purchased from Smolecule (San Antonio, TX, USA), and both ACh and choline standards were purchased from Sigma-Aldrich (Burlington, MA, USA). Betaine was supplied by Spectrum Chemicals (New Brunswick, NJ, USA) and levamisole was provided by MP biomedicals (Santa Ana, CA, USA). Fisher Scientific supplied all other chemicals.

Collection of pseudocoelomic fluid and identification of β-alanine betaine

We describe here observations that suggest that β-alanine betaine (Fig 1) may be an additional endogenous cholinergic ligand in Ascaris suum. The pseudocoelomic fluid of A. suum is a protein rich, metabolically active fluid that distributes metabolites and nutrients along the length of the nematode. This fluid is found between the intestine and cuticle of the worm bathing the muscle and nerves. ACh has already been found to be present in A. suum pseudocoelomic fluid [15]. To determine if other cholinergic transmitters are present in the pseudocoelomic fluid, we dissected adult female A. suum (N = 4) and drained the fluid from the parasite. We added 20 µM neostigmine to inhibit esterases that could degrade cholinergic compounds in the fluid.

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Fig 1. Chemical structures of acetylcholine, choline, betaine, β-alanine betaine, propionylcholine, butyrylcholine and butyrobetaine.

https://doi.org/10.1371/journal.pntd.0014518.g001

The fluid was analyzed using Liquid Chromatography-Mass Spectrometry (HILIC-MS). We observed signals consistent with ACh (S1 Fig) choline, and β-alanine betaine but did not observe evidence of betaine, propionylcholine, butyrylcholine or, butyrobetaine. ACh, synthesized from choline, is the principal excitatory neurotransmitter in nematodes and is extensively characterized through biochemical and electrophysiological studies. Choline has been identified previously in the nematodes C. elegans [16]; Necator americanus and Nippostrongylus brasiliensis [17]. Betaine has been identified previously in C. elegans [18]; H. contortus [19]; N. Americanus; and N. brasiliensis [17,20]. We confirmed with HILIC-MS/MS the presence of choline (S2 Fig) but did not find evidence of betaine in A. suum. Interestingly, we detected in the pseudocoelomic fluid of A. suum an unknown compound with a retention time of 3.39 min (Fig 2A). After further analysis, the compound was hypothesized to be β-alanine betaine (Fig 1).

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Fig 2. Chromatograms and HILIC–MS/MS showing signals for β-alanine betaine.

A: Chromatogram from A. suum pseudocoelomic fluid for suspected β-alanine betaine. B: Chromatogram for β-alanine betaine neat chemical standard. C: MS/MS spectrum for suspected β-alanine betaine from A. suum pseudocoelomic fluid. D: MS/MS spectrum for β-alanine betaine neat chemical standard.

https://doi.org/10.1371/journal.pntd.0014518.g002

To verify that the unknown compound was β-alanine betaine, a standard was purchased from Smolecule, and analysis with HILIC-MS/MS was performed. The β-alanine betaine standard matched the profile of the presumed β-alanine betaine from the pseudocoelomic fluid with an identical retention time of 3.39 min (Fig 2B), and Tandem Mass Spectrometry peaks at 58.06, 59.07, 60.08, 73.03, and 132.10 m/z (Fig 2C and 2D). Signals consistent with β-alanine betaine were observed in all four samples.

Pretzel effect of injected β-alanine betaine

With the detection of β-alanine betaine in the A. suum pseudocoelomic fluid, we investigated if this compound modulates motility like other cholinergic compounds. We injected either 500 µL of APF salt solution (negative control: N = 4), 500 µL 30 µM levamisole (positive control: N = 4), 500 µL 30 µM betaine (N = 4), or 500 µL 30 µM β-alanine betaine (N = 4), into separate free moving adult female A. suum using a fine needle, just in front of the gonopore and compared the effects on motility. The APF injection did not affect the continuous slow movement of any of the worms or their posture (Fig 3A). The injection of levamisole rapidly inhibited movement and all worms straightened into rod-like structures (Fig 3B). The injection of betaine, like APF, did not affect the slow movement or posture of the worms (Fig 3C). However, the effect of β-alanine betaine was different to the other compounds as all worms slowly became coiled and formed a ‘pretzel’ like structure (Fig 3D). These results demonstrate that β-alanine betaine promotes a unique paralytic phenotype in A. suum, potentially by acting on cholinergic receptors.

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Fig 3. Injection of β-alanine betaine promotes “pretzel” paralysis phenotype.

A: Adult female A. suum injected with 0.5mL APF, the worm continued to move with no clear effect, n = 4. B: Adult female A. suum injected with 0.5mL 30 µm levamisole: note the straight rod paralyzed appearance, n = 4. C: Adult female A. suum injected with 0.5mL 30 µm betaine: the worm continued to move with no striking effects, n = 4.D: Adult female A. suum injected with 0.5mL 30 µm β-alanine betaine. Note the coiling (pretzel) posture, n = 4.

https://doi.org/10.1371/journal.pntd.0014518.g003

DEG-3/DES-2 subfamily subunits, including ACR-23, are present in Ascaris intestine and body wall

The different effects of injected levamisole and β-alanine betaine on A. suum paralysis suggests the two compounds act at different sites. Levamisole activates nAChRs which are composed of UNC-29, UNC-63, ACR-8, UNC-38 subunits which are not part of the DEG-3/DES-2 subfamily [2,21]. In C. elegans, betaine is an agonist of either homomeric nAChRs composed of ACR-23 or ACR-20 [7,8,22], DEG-3/DES-2 heteromeric receptors [9], or the betaine-gated chloride channel LGC-41 [10], suggesting that β-alanine betaine could act on similar receptors in A. suum.

In C. elegans, ACR-23 is expressed in different tissues, including body wall muscles [23]. To determine if acr-23 is present in A. suum, we performed a blast search with Cel-acr-23 (F59B1.9) against the A. suum genome (PRJNA62057) and identified the gene AgR001_g108, as the Asu-acr-23 orthologue with 53.6% similarity in the protein sequence to Cel-ACR-23 (S3 Fig). We generated primers targeting Asu-acr-23 (Table 1) and screened for the presence in paired cDNA pools of intestine and body wall (the body wall includes the muscle and nerve cords). Asu-acr-23 is present in both cDNA pools (Fig 4 and S1 Raw gel). To ensure that we successfully targeted Asu-acr-23 we performed Sanger sequencing by comparing the full length acr-23 product obtained from our cDNA pool against the AgR001_g108 sequence found in the WormBase Parasite database. Our full-length Asu-acr-23 was 99.8% similar to the database (S4 Fig).

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Fig 4. RT-PCR analysis of Asu-acr-23, acr-20, des-2, deg-3 and lgc-41.

A: paired body wall. B: intestine from female A. suum. Each lane represents one of the genes for the intestine or body wall of an individual worm. Asu-gapdh from the intestine or body wall was used as a positive control (Cb/Ci). NC = negative control, no cDNA template present. M = FastRuler Middle Range DNA Ladder (ThermoFisher Scientific).

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We also identified and screened for the presence of A. suum orthologues of the other C. elegans betaine sensitive channels including, ACR-20, DES-2 and DEG-3 and the inhibitory betaine activated ion channel LGC-41 as possible β-alanine betaine targets (Tables 1–2). We detected the presence of all four subunits in both the intestinal and body wall cDNA pools of A. suum (Fig 4 and S1 Raw gel), with Asu-acr-23 having the brightest band visually in the body wall. These results suggest that targets composed of these genes may mediate the effects of β-alanine betaine in A. suum.

Intestine and muscle cells responses to β-alanine betaine

With the identification of potential receptors for β-alanine betaine, we investigated if application to either the intestine or muscle elicited a response. We first tested the effects of β-alanine betaine on the intestine of A. suum by measuring the Ca²⁺ signal. One mM β-alanine betaine induced large and prolonged increases in enterocyte intracellular Ca²⁺, with a mean amplitude of 31% ± 2.09% (N = 3) (Fig 5A and 5B). Interestingly, the increase in the Ca²⁺ signal did not occur immediately after applying β-alanine betaine but instead took 2 minutes before the Ca²⁺ started to increase (Fig 5A). The Ca²⁺ signal started to slowly decline after β-alanine betaine was washed out before returning to near baseline levels with 79% ± 8.1% of intestinal regions having an increase in fluorescence (Fig 5C), suggesting that the receptors were not localized to specific regions of the intestine. To test for viability, 10 mM CaCl₂ was applied and we observed robust increases in Ca²⁺ (54% ± 1.8%) in 100% of regions (Fig 5B and 5C).

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Fig 5. β-alanine betaine stimulates robust increases in Ca²⁺ in A. suum intestines.

A: Representative response to 1mM β-alanine betaine. Red lines indicate mean fluorescence; black dotted lines represent ±SEM. Grey Box indicates stimulus application. B: Total mean maximal amplitudes of Ca²⁺ fluorescence to 1mM β-alanine betaine (grey bar) and 10mM CaCl₂ (white bar). C: Mean percent of regions showing a Ca²⁺ response to 1mM β-alanine betaine (β-alanine betaine: grey bar) and 10mM CaCl₂ (white bar). n = 3 intestines from 3 individual female A. suum with 150 total regions. β-alanine betaine 119/150 regions responding, CaCl₂ = 150/150 regions responding. All values represented as means ± SEM.

https://doi.org/10.1371/journal.pntd.0014518.g005

We also measured the effects of β-alanine betaine on the membrane potential of A. suum body muscle cells. We applied 10 sec applications of 30 µM, 100 µM, 300 µM and 1 mM β-alanine betaine to body muscle cells and compared the effects to flanking 10 µM ACh applications (Fig 6). In each of the muscle cell preparations, the depolarization in response to β-alanine betaine was slower and weaker when compared to ACh but demonstrated a concentration dependent effect, with 30 µM β-alanine betaine having an mean ΔmV of 0.06 mV ± 0.03, 100 µM having a ΔmV of 0.4 ± 0.2, 0.3 mM having an mean ΔmV of 1.2 mV ± 0.3 and 1 mM having an mean of 2.2 mV ± 0.4 (Fig 6A and 6B). We observed no significant difference between the flanking 10 µM ACh applications (6.9 mV ± 1.1 and 7.1 mV ± 0.9) (Fig 6B). We also observed a concentration dependent effect on the rate of spontaneous depolarizations (spikes) that took more than a minute before reaching full effect with 1 mM β-alanine betaine having the highest frequency 79.3 spikes min^-1 (± 16.4) (Fig 6A and 6C). Again, we observed no significant difference between the flanking 10 µM ACh applications (25.5 ± 5.1 and 25.75 ± 4.1) (Fig 6C). These results demonstrate that β-alanine betaine has effects on A. suum muscle depolarization and intestinal Ca²⁺ signaling suggesting a novel functional cholinergic ligand in an anerobic parasite.

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Fig 6. β-alanine betaine stimulates small depolarizations on the muscle bags of A. suum.

A: An intracellular membrane potential recording from the muscle bag-region of a female A. suum worm to higher concentrations of β-alanine betaine. Samples were first exposed to an application of 10 µM ACh by rapid perfusion for 10sec followed by an application of 300 µM β-alanine betaine for 10 sec, 1 mM β-alanine betaine for 10 sec and finally to a second application of 10 µM ACh for 10 sec. B: Histograms of the mean + /- S.E.M. peak depolarization responses to the first 10 µM ACh application (grey bar), 30 µM β-alanine betaine, 100 µM β-alanine betaine (green bar), 300 µM β-alanine betaine (blue bar), 1 mM β-alanine betaine (red bar), and the second application of 10 µM ACh (grey bar; hashed). N.S. not significantly different to 1^st 10 µM ACh (1^st 10 µM ACh vs 2^nd ACh, P = 0.799, t = 0.2613, df = 11, paired t-test). C: Histograms of the mean + /- S.E.M. number of spikes produced during the application of first 10 µM ACh application (grey bar), 30 µM β-alanine betaine (pink bar), 100 µM β-alanine betaine (green bar), 300 µM β-alanine betaine (blue bar), 1 mM β-alanine betaine (red bar), and the second application of 10 µM ACh (grey bar; hashed). N.S. not significantly different to 1^st 10 µM ACh (1^st 10 µM ACh vs 2^nd ACh, P = 0.931, t = 0.088, df = 11, paired t-test). ACh recordings n = 12 individual muscles from 12 individual A. suum females. 300 µM and 1 mM β-alanine betaine n = 7 total recordings from 7 individual A. suum females. 100 µM β-alanine betaine n = 5 total recordings from 5 individual A. suum females. 30 µM β-alanine betaine n = 4 total recordings from 4 individual A. suum females. All values represented as means ± SEM.

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Effects of β-alanine betaine on DEG-3 C. elegans subfamily null mutants

With evidence that β-alanine betaine affects the movement and produces a pretzel shaped A. suum after injection, we utilized C. elegans to determine if an effect of β-alanine betaine was species specific. Following previous techniques with betaine and using established techniques with C. elegans with high concentrations that are required to cross their cuticle and to overcome bacterial metabolism [24,25], we incubated wild-type N2 C. elegans animals on E. coli OP50 seeded NGM plates containing 50 mM β-alanine betaine. The motility of C. elegans decreased progressively over 2 hours with 77% (± 3.3%) of N2 worms being paralyzed by the end of the experiment (Fig 7A: orange line). The N2’s lost their stereotypical sinusoidal wave phenotype (Fig 7B). With ACR-23 being responsible for betaine mediated signaling in C. elegans, we hypothesized that β-alanine betaine would act on similar nAChRs. We exposed acr-23(ok2804) nulls to 50 mM β-alanine betaine and observed reduced paralysis compared to N2, with only 19% (± 4.4%) of animals being paralyzed after 2 hours (Fig 7A: red line). Unlike N2, acr-23 nulls still maintained the sinusoidal waveform (Fig 7C). We repeated the experiments using 50 mM betaine and saw no evidence of larval arrest or animal paralysis like previous publications [7,10] (S5 Fig). These results suggest that β-alanine betaine paralyzes C. elegans with effects mediated via ACR-23 receptors.

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Fig 7. β-alanine betaine paralyzes C. elegans through DEG-3/DES-2 subfamily members.

A: Motility assay on 50 mM β-alanine betaine with wild-type N2 C. elegans (Orange), acr-23 (red), acr-20 (blue), deg-3/des-2 (green) and lgc-41 (pink). n = 30 worms for each genotype, 3 replicates. All values represented as means ± SEM. B: Photograph of N2 after 2 hours on 50 mM β-alanine betaine. Note the lack of sinusoidal wave. C: Photograph of acr-23 after 2 hours on 50 mM β-alanine betaine. Note the presence of body bends in the worm.

https://doi.org/10.1371/journal.pntd.0014518.g007

We then treated other C. elegans null mutants of the betaine receptors on our β-alanine betaine plates. Fig 7A shows the effects of 50 mM β-alanine betaine on acr-20 (ok1849), deg-3 (u662)/des-2 (u695) double mutants and lgc-41(sy1495) null mutants. Over the 2-hour period, β-alanine betaine affected the nulls differently: the most resistant worms were the acr-20 null mutants, (Fig 7A: blue line). Only 25% (± 3.3) of the acr-20 nulls were paralyzed after 2 hours, a result like the acr-23 null mutants. Both the acr-20 nulls and acr-23 nulls were still capable of body bends. The DEG-3/DES-2 (u662/u695) double mutant was less resistant to β-alanine betaine compared to acr-23 and acr-20, with 35% (± 5.8%) of worms being paralyzed after two hours (Fig 7A: green line). The lgc-41 mutants were even less resistant with 45% (± 5.8%) of worms being paralyzed (Fig 7A: pink line), after 2 hours. Our results suggest that β-alanine betaine could act on the ACR-23 and ACR-20 receptors but also less potently on the DEG-3/DES-2 heteromeric receptors and LGC-41 receptors.

Betaine in aerobic and β-alanine betaine in anaerobic environments

Betaine may be produced by oxidation of choline or be taken up from the environment: in C. elegans choline is first oxidized to betaine aldehyde by choline dehydrogenase and then to betaine, by betaine aldehyde dehydrogenase [7]. Betaine can also be taken up from the environment by a high-affinity transporter SNF-3 [7]. In parasitic nematodes, betaine has been found to be present in H. contortus, N. americanus and N. brasiliensis [17,19,20].

We found β-alanine betaine but no betaine in adult A. suum which live in a low oxygen environment in the host pig intestine. In hypoxic plants [26], β-alanine betaine is synthesized from β-alanine via methyltransferases: β-alanine is methylated to N-methyl β-alanine, then methylated again to N-N-dimethyl β-alanine and then again to β-alanine betaine. This pathway relies exclusively on methyltransferases and is independent of oxygen, in contrast to choline oxidation pathways required for betaine synthesis. β-alanine betaine is preferentially accumulated in organisms adapted to high saline and hypoxic or anaerobic conditions [27]. Thus, it is possible that A. suum in its anaerobic environment produces β-alanine betaine rather than betaine as an endogenous ligand.

Ion-channels activated by betaine and choline in C. elegans and H. contortus

Choline and betaine have been reported in C. elegans [22,28]. Both choline and betaine serve different functions, including acting as osmolytes, and as endogenous ligands of nAChRs. Choline and betaine selectively activate nematode members of the DEG-3/DES-2 subfamily of nAChR ion-channels. Expressed C. elegans ACR-23 homomeric nAChR channels are activated by betaine (EC₅₀ = 1.4 mM) but not choline or ACh [7]. C. elegans ACR-20 homomeric nAChR channels are selectively activated by betaine (EC₅₀ ~ 25 µM), less by choline (EC₅₀ ~ 1.2 mM) and hardly at all by ACh [8]. C. elegans DEG-3/DES-2 channels are activated by betaine (EC₅₀ ~ 0.6 mM) and less potently by choline (EC₅₀ ~ 1.8 mM) [9]. Betaine is more potent than choline and ACh on these C. elegans channels. β-alanine betaine has not yet been tested on expressed nAChR channels.

Betaine is also found in H. contortus [19]. Studies of H. contortus expressed MPTL-1 channels (formally referred to as Hc-ACR-23-H) found that betaine (EC₅₀ = 41 µM) activates these channels, and that choline (EC₅₀ = 1.3 mM) is less potent [8]. Rufener et al. [29] studied effects of ACh and choline on expressed H. contortus DEG-3/DES-2 channels and found that choline had an EC₅₀ of 10 mM, but that ACh was a poor agonist even at 10 mM; the effects of betaine on this nAChR were not studied.

Although we found signals for ACh, choline, and β-alanine betaine in pseudocoelomic fluid of A. suum, we did not find betaine. Betaine is produced by oxidation of choline and requires an aerobic environment; the absence of betaine and the presence of β-alanine betaine in A. suum may be explained by its anaerobic environment and that β-alanine betaine is produced anaerobically [26,27]. Thus, if betaine functions as both an osmolyte and an endogenous ligand in aerobic nematodes, β-alanine betaine may fulfill analogous roles in anaerobic species such as A. suum.

Effects of monepantel illustrate the therapeutic significance of the DEG-3/DES-2 subfamily

Monepantel (an ADD: amino-acetonitrile derivative) is active against nematode parasites of ruminants including H. contortus, Trichostrongylus colubriformis, Nematodirus spathiger, Cooperia oncophora and Teladorsagia circumcincta. Monepantel also inhibits movement of C. elegans by causing hypercontraction of the body wall and anterior pharynx. This hypercontraction is mediated in part by acr-23 nAChRs because C. elegans null-mutants are resistant to ADDs [6]. Furthermore, monepantel is a type 2 positive allosteric modulator (PAM) on members of the DEG-3/DES-2 subfamily [7]. Combinations of monepantel and betaine positively modulate signals in C. elegans via ACR-23, ACR-20 and in the H. contortus channel MPTL-1 [7], but has no PAM effect on betaine signaling on the DEG-3/DES-2 channels [9]. Monepantel is not effective against A. suum infections, but is a non-competitive antagonist of ACR-16’s and on the levamisole and pyrantel nAChRs [30]. Even high concentrations (10 µM) of monepantel do not produce contraction of A. suum body flaps.

β-alanine betaine as an endogenous ligand in A. suum

Betaine activates members of the DEG-3/DES-2 subfamily in aerobic nematodes. In the absence of detected betaine, we suggest that β-alanine betaine may substitute functionally for betaine in A. suum to activate members of the DEG-3/DES-2 subfamily. This view is supported by: 1) the absence of betaine and presence of β-alanine betaine in the pseudocoelomic fluid in A. suum; 2) the absence of an effect of betaine and the pretzel effects of β-alanine betaine following injection in A. suum; 3) the increase in cytoplasmic Ca²⁺ in enterocytes following application of β-alanine betaine; 4) the depolarizing effects of β-alanine betaine on A. suum somatic muscle cells; and 5) the presence of Asu-acr-20, Asu-acr-23, Asu-deg-3, Asu-des-2, and Asu-lgc41 in the body wall and intestine enterocytes. We point out however that the aerobic C. elegans null mutants of acr-20 (ok1849), acr-23 (ok 2804), deg-3 (u662)/des-2 (u695) and lgc-41(sy1494) were resistant to β-alanine betaine which suggests that the DEG-3/DES-2 subfamily in C. elegans is also sensitive to β-alanine betaine.

Conclusion

Nematodes have multiple subfamilies of nAChRs. We have identified β-alanine betaine in addition to ACh and choline in A. suum. We found evidence that β-alanine betaine can act on the DEG-3/DES-2 subfamily of nAChRs. Multiple subfamilies of nAChRs and multiple endogenous cholinergic ligands permit an increase in behavioral complexity, including movement. Given that the DEG-3/DES-2 subfamily of nAChRs is not present in mammals, it also provides opportunities for the development of selective anthelmintics.

Financial disclosure

This study was supported by NIH NIAID Grants R01AI047194, R01AI155413, to RJM, the EA Benbrook Endowed Chair of Pathology and Parasitology and by ISU CVM Seed Grant funding from the Dr Stephen G. Juelsgaard Dean’s award to PDW from Iowa State College of Veterinary Medicine. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. PDW received salary from the R01AI155413, NIH NIAID Grant.