Ascaris Mouse Model Protocols: Advancing Research on Larval Ascariasis Biology

Abstract

Ascariasis, caused by both Ascaris lumbricoides and Ascaris suum, is the most prevalent parasitic disease worldwide, affecting both human and porcine populations. However, due to the difficulties in assessing the early events of infection in humans, most of the studies on human ascariasis are restricted to the chronic intestinal phase. Therefore, developing an Ascaris mouse model has become a fundamental tool for investigating the immunobiology and pathogenesis of the early infection referred to as larval ascariasis due to its practicality and ability to replicate the natural processes involved. The Ascaris mouse model has been widely used to explore factors such as infection resistance/susceptibility, liver inflammation, lung immune-mediated pathology, co-infections, and notably, as a pivotal element in pre-clinical vaccine trials. Exploring the immunobiology of larval ascariasis may offer new insights into disease development and provide a substantial understanding of key components that trigger a protective immune response. This manuscript focuses on creating a comprehensive guide for conducting Ascaris experimental infections in the laboratory as a foundation for future research efforts.

Basic Protocol 1:

Obtaining and embryonating Ascaris suum eggs from adult females

Alternate Protocol 1:

Cleaning and purifying Ascaris suum from female’s uteri

Basic Protocol 2:

Ascaris suum egg preparation and murine infection

Basic Protocol 3:

Measuring larval burden and Ascaris larval-induced pathogenesis

Basic Protocol 4:

In vitro Ascaris L3 larvae hatching and purification

Support Protocol 1:

Crude antigen preparation from Ascaris infectious stages

Basic Protocol 5:

Ultrastructure-expansion microscopy (U-ExM) of Ascaris suum larval stages.

INTRODUCTION:

Ascariasis, caused by both Ascaris lumbricoides and Ascaris suum (Wang & Davis, 2020), is one of the most common parasitic diseases worldwide, affecting both humans and pigs. It is considered a major global health problem for humans (Holland et al., 2022; Hotez et al., 2006) and poses significant economic losses for the pig farming industry (Knecht et al., 2012). The disease is endemic in more than 150 countries, with recent estimates suggesting that approximately 732 million people are infected with around 2,090 deaths annually (Holland et al., 2022; IHME, 2019). While Ascaris suum has traditionally been linked with porcine infection, numerous epidemiological and experimental studies have indicated its significant role as a zoonotic agent in human infections (Arizono et al., 2010; Nejsum et al., 2012; Nejsum et al., 2005; Silva et al., 2021; Takata, 1951).

The clinical manifestation of the disease in the natural host can be divided into two distinct phases: the acute or larval ascariasis, characterized by a robust inflammatory response due to a mandatory transient migration of larval stages in different host tissues, including gastrointestinal (GI) tract, liver, lungs, and airways; and the chronic intestinal phase, associated with the establishment and maturation of adult worms in the intestinal lumen with oviposition of thousands of eggs daily (Crompton, 2001; Dold & Holland, 2011; Sinniah, 1982). Diagnostic techniques largely still rely on the presence of Ascaris eggs in the stools usually by the identification of eggs microscopically, though molecular-based diagnostics have been used as well. These diagnostic approaches might explain why most studies on human ascariasis are restricted to the chronic intestinal phase. However, to understand more comprehensively Ascaris parasites and parasitism, it is essential to conduct new studies focusing on the immunobiological aspects of the early Ascaris spp. infection. To address these knowledge gaps, including the investigation of the immunological underpinnings of the larval ascariasis, researchers have utilized several animal models involving Ascaris spp. over the years, including pigs, mice, guinea-pigs, rabbits, lambs, goats, gerbils, rats and cows [(reviewed (Holland, 2021)]. While pigs naturally serve as hosts and effective models for both larval ascariasis and chronic infection, their utility as an experimental model is restricted by the logistical and financial difficulties associated with handling and housing larger animals (Holland, 2021). As a result, the mouse model has become the leading in vivo method for immunoparasitological investigations, owing to its practicality and its ability to replicate the biological processes and pathways observed during the early stages of infection in the definitive host (Gazzinelli-Guimaraes et al., 2013; Lewis et al., 2006; Mitchell & Lewers, 1976; Murrell et al., 1997; Slotved et al., 1998).

Indeed, the Ascaris mouse model (Figure 1) has been widely utilized for studies of liver inflammation (Deslyper et al., 2019; F. M. S. Oliveira et al., 2022), type 2 immune responses in the lung tissue (Gazzinelli-Guimaraes et al., 2019; Gazzinelli-Guimaraes et al., 2013; Weatherhead et al., 2018), host genetics underlying susceptibility to infection (Dold et al., 2010; Lewis et al., 2006; L. M. Oliveira et al., 2022), re-infections (Nogueira et al., 2016), co-infections and interactions (Gazzinelli-Guimaraes et al., 2017; Gazzinelli-Guimaraes et al., 2019; Oliveira et al., 2019; Vieira-Santos et al., 2021), and finally, as an important tool for pre-clinical vaccine trials (Castro et al., 2023; Gazzinelli-Guimaraes et al., 2018; A. C. Gazzinelli-Guimaraes et al., 2021; Wei et al., 2017), because it allows for the evaluation of infection-driven pathogenesis as well as vaccine-driven immunogenicity at a tissue level.

Figure 1: Ascaris mouse model and the associated pathogenesis of larval ascariasis.

Fully embryonated and decorticated eggs are introduced through oral gavage using intragastric intubation, targeting the stomach. During the initial hours of infection, the infective L3 larval stages hatch from the eggs within the small intestine. They subsequently migrate to the large intestine, where they breach the cecal and colonic walls, entering the bloodstream, and initiating the hepatic-tracheal migration phase. The disruption of the large intestinal mucosa by these migrating larval stages facilitates bacterial translocation, the implications of which for pathogenesis are still relatively understudied. Once within the vascular system, carried by the portal circulation, the larvae make their way to the liver. The peak of infection in the liver typically occurs between day 1 and day 5 post-infection, depending on the specific mouse strain. Notably, in natural hosts such as pigs, a highly infected liver often exhibits distinct macroscopic “white spots” as the result of a robust inflammatory infiltration in the liver parenchyma, a process that leads to tissue necrosis and functional impairment of the organ. Subsequently, using pulmonary circulation, L3-liver stages migrate to the lungs, peaking in pulmonary tissue on days 7–8 following infection. Upon reaching the lungs, these larvae traverse the lung parenchyma and penetrate the alveolar spaces, resulting in hemorrhage and mechanical damage within the organ. Macroscopic hemorrhagic spots in Ascaris-infected lungs are a common visual observation (see also Figure 15). Immunologically, lung-stage larval migration provokes an early influx of neutrophils correlated with levels of IL-6, followed by a dominated eosinophil-dominated type 2 immune response (See Commentary section). However, during pulmonary migration in naive mice, this robust type 2 response is only fully mounted when most of the larvae have reached the airways, allowing for the progression of the life cycle in mice and the establishment of a chronic and persistent infection in the natural host. By days 8–10 post-infection, the L3-lung stages migrate to the airways, as part of their journey back to the intestine, a process generally completed by days 10–12. Upon returning to the small intestine, the larval stages undergo a molt thereby becoming L4s by days 12–14 post-infection. The arrival in the gut induces a dysbiotic microbiome characterized by shifts in composition and abundance. Typically, Ascaris-infected mice clear the infection around day 14.

Developing a standard Ascaris mouse model is of fundamental importance to understanding the biology of early events of Ascaris infection, which remains not fully understood. Therefore, our aim was to standardize common guidelines for murine infection with Ascaris parasites. Here, we provide a series of protocols that have been successfully employed in several laboratories worldwide to understand the biology of the initial events elicited by Ascaris infection. This includes guidelines for isolating parasite eggs from adult females, embryonating them in vitro (refer to Basic Protocol 1 and Alternate Protocol 1), and preparing them for mouse infection, as well as elucidating the infection process itself (refer to Basic Protocol 2). We also explore detailing methods to measure parasite burden and the resultant pathogenesis in mice (refer to Basic Protocol 3). Additionally, we discuss the recovery of Ascaris larval stages from eggs and infected mouse tissues, which holds multifaceted potential for various research methodologies, including antigen preparation, as well as ultrastructure-expansion microscopy (U-ExM) using different parasite larval stages (refer to Basic Protocols 4 and 5).

Of note, all the protocols outlined in this guideline are applicable to both Ascaris suum and Ascaris lumbricoides species. However, the most efficient way to obtain Ascaris eggs is by isolating them directly from the uterus of adult females. Typically, Ascaris. lumbricoides adult worms are expelled naturally in the feces of infected individuals residing in endemic areas, or after anti-helminthic chemotherapy; and Ascaris suum adult worms are normally obtained from infected pigs in slaughterhouses or farms. In this way, due to the easier access to Ascaris suum parasites, this species has been widely utilized for the Ascaris mouse model.

Safety

CAUTION: Ascaris sp. is a Biosafety Level [2 ([BSL-2]) pathogen. Follow all appropriate guidelines and regulations for the use and handling of pathogenic microorganisms. Moreover, once fully embryonated, both Ascaris suum and Ascaris lumbricoides eggs become infective to humans. Personnel should wear laboratory coats, gloves and masks when working with this biohazard parasite. Decontaminate work surfaces after use and after spills with standard agents including 70% ethanol, 1% hypochlorite, or soap solutions. Any material contaminated with Ascaris eggs should be either boiled for at least 30 minutes or discarded in medical pathological waste (MPW) boxes for incineration.

Animal Facility Containment

Ascaris sp. larvae do not develop into adult worms in mice, which means that viable eggs will not be produced in the stools of the infected mice. Because of that, Ascaris are not transmitted to other mice in a facility and cross-transmission is not seen even when infected mice are mixed with uninfected mice in the same cage.

BASIC PROTOCOL 1:

OBTAINING AND EMBRYONATING ASCARIS SUUM EGGS FROM ADULT FEMALES

One of the challenges of working with the Ascaris mouse model in the laboratory is the fact that Ascaris lumbricoides and Ascaris suum are fully permissive parasites only to their definitive hosts, which include humans and pigs. Thus, the infection in mice which clears itself before the establishment of adults in the intestine, will not produce viable eggs for sustained and cyclic parasite maintenance in the lab. In contrast to other murine models for helminth parasites, including Heligmosomoides polygyrus, and Trichuris muris, which are natural rodent nematodes, working with Ascaris in the lab will always require a new source of embryonated eggs for new infections in mice.

This first protocol is a compilation of protocols from different laboratories which provides important details in obtaining, purifying and embryonating Ascaris suum eggs from adult female uteri.

Materials

Adult Ascaris suum worms - sourced from infected pigs, acquired either from carcasses in slaughterhouses or through experimental infection.

Phosphate-buffered saline – PBS (pH 7–7.5)

0.2 M Sulfuric acid

100 mL plastic or glass beakers

15 cm plastic or glass petri/culture dishes

Surgical scissors and tweezers

Pestle and mortar

100 μm strainer for 50mL Tubes [Filter material: Polyethylene terephthalate (Greiner, Bio-One, Cat.# 542000]

50 mL conical tubes

Benchtop centrifuge with 50mL tubes’ adapter

75 cm² flasks with filters

Incubator at 26°C

Light microscope

Protocol steps with step annotations

1. Transfer all recovered Ascaris suum adult worms to beakers and manually rinse them with PBS five times.

2. In 15 cm Petri/culture dishes, separate the adult females (usually longer with their tails in a straight position) from the males [see Figure 2 (right-A) for macroscopic differences].

3. Using only the adult females, cut at genital girdle or just posterior (the constriction encircling the anterior third of body where the vulva is located) at the location where the gonads end or just posterior to this (look inside worms for splitting of the thick ends of the uterus and ending of gonad). Note that the anterior end is thinner [see Figure 3 (especially 3A) for guidance].

4. The pair of uteri should pop out of the cut region of the worm (if they do not, try to gently squeeze posterior to cut and run fingers anterior to end to push uterus out). Note that if there is not a pair of tubes and no thickening of the pair in the anterior of the worm that appears to exit via a small duct, that you probably have a male.

5. Gently pull uteri out and up into the air (they should easily leave the body unless the tissue has substantially decayed). If the worms are relatively fresh, the spaghetti-like material of the oviduct and ovary and the flat, brown intestine should remain inside the worm.

   NOTE: If not, you may remove any oviduct, ovary, or intestine that clings to the upper half of the uteri, although it is not necessary.

6. Examine the length of the uterus cut 1/4 of the way down. Collect the proximal 1/4 (i.e., the region that is thickest and closest to the vulva) and transfer into 20 mL of PBS in a large beaker on ice. The length of the cut region will vary since the lengths of worms and uteri are quite variable.

   NOTE: Depending on the amount of material you collect, you can use less or more PBS; just make sure to keep the material submerged.

7. Transfer all collected uteri to a beaker and rinse pooled uteri 3x with PBS.

   NOTE: Following step 7, you have the option to expedite the purification process by proceeding directly to step 8. Alternatively, for a more thorough and debris-free purification, we recommend to follow Support Protocol 1.

8. Isolate the eggs from all uteri by mechanical maceration using a pestle and mortar with PBS (Figures 3C and D).

9. Transfer all the homogenates to a new 50 mL tube and fill with PBS to 50 mL.

10. Purify the macerate by filtration on 100 μm strainers attached to new 50mL tubes (Figure 3E).

11. Centrifuge the eggs at 1000 relative centrifugal force (rcf) for 10 min at room temperature (RT) and resuspend in 50 mL PBS-0.1 M sulfuric acid (H₂SO₄).

12. Place 25 mL of the eggs in 75 cm² culture bottles, at a concentration of 25 eggs/μL (higher egg density inhibits embryonation) in incubators at 26°C for a maximum of 200 days.

    NOTE: The peak of Ascaris egg infectivity for experimental infections varies between 100–150 days in culture, however, the eggs become infective around day 35.

    NOTE: For egg counting, use three aliquots of 10 μl each and average the number of eggs in this volume. Then, use a proportion to determine the volume required to achieve a concentration of 25 eggs/μl.

13. During incubation, manually agitate the bottles once a week.

14. Check the egg embryonation by assessing the larval development inside the eggs (Figure 4) and calculate the embryonation rate of the fully embryonated eggs under the microscope every 10 days (analysis of three independent counts of 10 μL of egg suspension).

    NOTE: Typically, Ascaris suum embryonation rates vary between 50% and 95% following 6–8 weeks of embryonation due to some unfertilized eggs and eggs that remain undeveloped in the egg suspension.

Figure 2: Ascaris suum adult worms.

Females are usually longer and more robust than the males and present their tail in straight position (A), in contrast to the males with the tails curved ventrally (B).

Figure 3: Dissection of Ascaris suum adult female for egg extraction.

(A) Female adult worm dissected longitudinally to expose the internal organs. The arrow indicates the position of the uterus, located closest to the vulva. (B) The uterus is carefully removed and transferred to a mortar, for its maceration with a pestle (C) until a homogeneous solution is observed (D). Subsequently, the solution is filtered in a 100 μm strainer to purify eggs for further use (E).

Figure 4: Ascaris suum eggs embryonation.

Embryonation progression of Ascaris suum eggs from day 0 (non-embryonated) to day 40 (fully embryonated containing the L3 larval stages) of culture at 26°C with 0.1 M H₂SO₄.

ALTERNATE PROTOCOL 1

CLEANING AND PURIFYING ASCARIS SUUM EGGS FROM FEMALE’S UTERI.

The mechanical maceration of the females’ uteri to isolate eggs results in a heterogeneous suspension of the eggs contaminated with debris from uteri tissues. The filtration of the eggs in a 100 μm strainer improves the purity of the egg suspension but it is still possible to observe debris when they are examined microscopically. Of note, the maximum efficiency of embryonation of the eggs might be affected by contaminating debris. Below we provide an alternate method to clean any contaminating debris from the egg suspensions by multiple rounds of NaOH incubation. This method also removes the egg outer mamillated and albuminous layer, which is not needed for the development of the eggs.

Materials

Ascaris’s adult female uterus

Phosphate-buffered saline - PBS

0.5 N NaOH

0.1 M Sulfuric acid

Ice-cold MilliQ water

50 mL conical tubes

250 mL conical tubes

75 cm² flasks with filters

Incubator at 26°C

Heating Magnetic Stirrer

Optical microscope

Benchtop centrifuge with 50mL tubes’ adapter

Beaker

Protocol steps with step annotations

1. Start this protocol by measuring the uterus volume in a beaker after step 7 from Basic Protocol 1.

2. Treat the uteri with 0.5 N NaOH for ~1 hour while stirring at RT.

   NOTE: A typical ratio of uteri/NaOH is 250 mL packed uteri/3 L of 0.5 N NaOH.

3. Allow to settle ~ 45 min.

4. Decant the liquid carefully until ~400 mL or less remains in the beaker.

5. Treat again for 1 hour with 0.5 N NaOH (same proportion).

6. Allow to settle ~45 min.

7. Carefully decant the liquid until approximately 200–300 mL remains in the beaker, taking care not to disturb the settled eggs.

8. Spin the settled eggs in 250 mL conical bottles (filled to the top with cold MilliQ water) at 1250 rcf for 15 min.

9. Discard the supernatant and wash the pelleted eggs 3x by resuspending them using ice-cold MilliQ water (1250 rcf for 15 min).

10. Resuspend the eggs in 50 mL of PBS-0.1 M sulfuric acid (H₂SO₄).

    NOTE: Alternatively, the eggs can be resuspended in PBS pH=2 and stored at 4°C for 5 years or more.

11. Place 25 mL of the eggs in 75 cm² culture bottles, at a concentration of 25 eggs/μL, and bottles in incubators at 26°C for a maximum of 200 days. Of note, Ascaris eggs can also be incubated at 30°C. At this temperature, it takes about 10 days to reach to L1 and about 21 days to reach to L3 infectious stage (Wang & Davis, 2020).

BASIC PROTOCOL 2:

ASCARIS SUUM EGG PREPARATION AND MURINE INFECTION

In nature, Ascaris eggs may appear in three different forms: unfertilized, fertilized corticated and fertilized decorticated. Fertilized corticated eggs are round-shaped, 45–75 μm in diameter and have a thick shell with an external mammillated layer. In some cases, the outer layer is absent (decorticated eggs). Of note, Ascaris eggs isolated directly from the female’s uteri, are decorticated eggs (Figure 4). Regarding their ultrastructure, Ascaris sp. eggs are composed of a 3- to 4-μm thick, four-layer shell, which plays a crucial role in protecting the developing embryo from harsh environmental conditions and providing a barrier against external factors (Mkandawire et al., 2022). Ascaris four-layer egg shells are formed by an inner lipoprotein layer, that consists of a unique mixture of 25% protein and 75% lipid-containing ascarosides and is responsible for the impermeability of the shell (Brownell & Nelson, 2006); a chitin/protein layer that provides structural strength, a vitelline layer, and an outer acid mucopolysaccharide/protein uterine layer. The outer three layers can be removed by soaking the eggs in a solution of hypochlorite, leaving only the inner lipoprotein layer (Barrett, 1976). Thus, to facilitate the larval hatching process during infection, many laboratories have utilized the protocol to treat the eggs with 5% sodium hypochlorite at 37°C and 5% CO₂ incubator, as follows:

Materials

Ascaris suum fully embryonated eggs (see Basic Protocol 1)

Males or females (6–8 weeks old) of any mouse strain, mostly commonly used C57BL/6, BALB/c and CBA mice (see Understanding Results section below)

Filtered water (don’t use distilled water)

Phosphate-buffered saline - PBS

5% sodium hypochlorite

Stock Flasks containing eggs prepared on Protocol 1

1 mL disposable syringes and oral gavage needles (24G x 25mm with a 2mm tip diameter curved or straight)

Serological pipettes

Serological pipette controller (also known as pipette gun)

50 mL conical tubes

Benchtop centrifuge with 50mL tubes’ adapter

Incubator at 37°C and 5% CO₂

Light microscope

Protocol steps with annotations

1. Calculate the number of Ascaris eggs needed for the experiment.

   2,500 fully embryonated eggs per mouse (always calculate an extra 30% of eggs needed since you lose a lot of eggs during washes).

2. To prepare the eggs for infection, use the stock flasks from Basic Protocol 1. Pipette three independent 10 μL aliquots and count only the fully embryonated eggs by light microscopy.

3. Aliquot the required volume into a 50 mL tube and centrifuge at 1000 rcf for 10 min at RT.

4. Remove all the supernatant with a serological pipette (do not invert the tube since the pellet containing the eggs is very delicate).

5. Resuspend the pellet by adding 5% sodium hypochlorite up to 25 mL (mix well).

   NOTE: Usually, the sodium hypochlorite is sold at 10%, so just use 20 mL and add 20 mL of filtered water to make a 5% solution.

6. Incubate the solution in a 37°C and 5% CO₂ incubator for 90 minutes (with the lid of the 50 mL tube partially open).

7. After incubation, raise the volume of each tube to 45 mL with filtered water (this and the next washing steps are performed to remove the hypochlorite from the solution).

8. Centrifuge 1000 rcf for 10 min at RT.

9. Remove all the supernatant with a serological pipette (do not disturb the egg pellet).

10. Repeat steps 7 and 8.

11. Remove the supernatant with a serological pipette and resuspend the pellet in 1 mL of PBS.

12. Count the embryonated eggs from three independent 10 μL aliquots using a microscope.

13. After counting, adjust the volume of egg suspension to obtain 2,500 eggs in 200 μL of PBS per animal.

    NOTE: Prepare a suspension of 2,500 eggs/200 μL/mouse. Multiply this proportion by the number of mice. Eg. 20 mice – 50,000 eggs/4 mL (always calculate an extra 20% of eggs needed since you lose some volume in the seringes during the infection).

14. Each mouse gets infected via the intra-gastric route with a gavage needle attached to a 1 mL syringe containing 2,500 fully embryonated eggs (see Basic Protocol 1) in 200 μL of filtered water or PBS.

    NOTE: Take special care to mix egg suspension immediately before each drawing of the syringe, as the embryonated eggs settle very quickly. Do one mouse at a time.

BASIC PROTOCOL 3:

MEASURING LARVAL BURDEN AND ASCARIS LARVAE-INDUCED PATHOGENESIS

The parasite burden and pattern of larval migration in the murine host have been well-studied over the last few years. Several authors have shown that there is a great disparity in susceptibility to the infection by the mouse strain [C57BL/6 shows higher susceptibility and CBA more resistant (See Figure 5 below – reuse from Dold et al. (Dold et al., 2010)]. Moreover, mouse age also plays an important role in parasite burden, as older mice (16 weeks old) have been demonstrated to be considerably more resistant than 8-week-old mice (Gazzinelli-Guimaraes et al., 2013).

Figure 5. Ascaris larval migration in different mouse strains.

Left: re-used with permission from Dold et al. (Dold et al., 2010) (License Number 5600430964691, this graph shows the changes in the mean larval burden in the liver and lungs of Ascaris-infected C57BL/6j and CBA/Ca mice. Right: re-used with permission from Gazzinelli-Guimaraes et al. (Gazzinelli-Guimaraes et al., 2013) (License Number 5600360191764, this panel shows the pattern of Ascaris suum larval migration in BALB/c mice in the large intestine (A), liver (B), lung (C) and small intestine (D), over 14 days of infection.

Materials

Ascaris-infected mice

Phosphate-buffered saline – PBS at 4°C

10% buffered formalin

CO₂ chambers

Sterilized scissors and tweezers

1 mL and 10 mL disposable syringes

15 mL and 50 mL conical tubes

Rack for 50 mL tubes

Plastic Petri-dishes

Plastic 3 mL Transfer pipette

Serological pipette

100 μm, 70 μm and 20 μm strainers for 50 mL conical tubes

Catheter (18 GA x 1.16 IN – BD Insyte^™Autoguard^™)

12-well plate

Benchtop centrifuge with 50mL tubes’ adapter

Incubator at 37°C

Inverted light microscope

Microscopy slide

Protocol steps with step annotations

• 1Perform the euthanasia of the Ascaris-infected mice in CO₂ chambers, at the selected time point according to the objective of each experiment (See Figure 5).
• 2Collect the bronchoalveolar lavage (BAL) fluid, liver, lungs and intestines for the analysis of the parasite burden as follows:

Parasite burden in the Bronchoalveolar Lavage (BAL)

Position the mouse on its back and secure it in a supine position [Refer to Figure 6 for guidance or refer to (Kalidhindi et al., 2021)].

Figure 6. Bronchoalveolar Lavage Fluid collection.

The diagram illustrates the procedure for Bronchoalveolar Lavage Fluid (BALF) collection from murine lungs. The process involves the insertion of a catheter into the trachea followed by instillation and aspiration of 1mL of PBS, twice. BALF, containing cellular and soluble components from the lung’s alveolar spaces, is then collected for analysis. This diagram was created on biorender.com.

• 3Using scissors and tweezers, make a small incision in the skin at the neck of the mouse, and carefully open the upper chest cavity to expose the trachea.
• 4Carefully dissect the trachea and expose a segment of the upper part.
• 5Prepare two syringes with 1 mL of cold PBS each.
• 6Insert a catheter into the trachea through a small incision to ensure proper fixation. Remove the needle and attach the first syringe.
• 7Gently inject 1 mL of cold PBS into the trachea and then slowly aspirate it back into the syringe to obtain the first wash.
• 8Pass the collected 1 mL BAL fluid through a 20 μm strainer attached to a 50 mL tube.
• 9Repeat the process with the second syringe of 1 mL of cold PBS to obtain the second wash.
• 10Pass it again through the same 20 μm strainer attached to a 50 mL tube.
• 11The lung-stage larval stages from the BAL fluid will be retained on the top of the strainer.
• 12Remove the 20 μm strainer from the tube and invert it carefully into a 12-well plate.
• 13With a transfer pipette, flush the strainer from the back with 2 mL of PBS to dispense all collected larvae into the bottom of the 12-well plate.
• 14Using an inverted light microscope count the number of larvae in each well.
• 15For the immunological characterization of the BAL, including cytokine and cellular profiling, centrifuge the 2 mL of BAL that flowed through the strainer (step 11) at 600 rcf for 10 minutes at 4 °C to separate the cellular components from the supernatant.
• 16Carefully collect the supernatant without disturbing the pellet and divide it into two aliquots of 1 mL each. Store the aliquots in labeled tubes and freeze them at −80°C for further analysis.
• 17The cells from the BAL can be used for further analysis.

Parasite burden in the liver, lung tissue and small intestine:

• 18Transfer the collected livers, lungs, and small intestines to individual plastic petri dishes and add one drop of PBS with a transfer pipette over each tissue.
• 19Assemble modified-Baermann apparatus by attaching a 100 μm strainer to a 50 mL tube (See Figure 7). Fill the tube with 30 mL of PBS.
• 20In the Petri dishes, chop the liver or lung tissues into very small pieces with surgical curved scissors.
• 21Using a 3 mL transfer pipette, transfer the minced tissue pieces into a separate strainer apparatus, ensuring all pieces are collected.
• 22Fill up the 50 mL tube with PBS ensuring all pieces are submerged in the PBS solution on the top of the strainer. Avoid bubbles (See Figure 7).
• 23The larval burden in the intestine is perform in two steps: Using a 10 mL syringe, wash the small intestine lumen with 10 mL of PBS and collect the intestinal lavage individually from the petri dishe to new 15 mL tube.
• 24With surgical scissors, open the small intestines longitudinally. Place the opened small intestine in a separate strainer apparatus and fill the tube with PBS, as in step 23.
• 25Place the rack with all strainer apparatus in an incubator set to 37°C and incubate the tissues for at least 5 hours (or overnight) to allow larval migration from the tissues and subsequent settling to the bottom of the tube.

  NOTE: Ascaris larvae tend to migrate from the tissue into the warm PBS. Therefore, make sure all tissues are very well chopped to improve larvae recovery.

• 26Carefully remove the rack from the incubator. Discard the strainers with the tissues.
• 27Collect the precipitate from the bottom of the tube using a transfer pipette and transfer it to a 15 mL tube.
• 28Centrifuge at 1000 rcf for 10 minutes at RT, including the tube with the intestinal lumen lavage.
• 29Discard the supernatant (do not disturb the pellet) using a 10 mL serological pipette then resuspend the pellet with either 1 mL of PBS or 1 mL of 10% buffered formalin. Adjust the volume based on the size of the pellet.

  NOTE: If the goal of the experiment is just to quantify the parasite burden, fix the larvae in 10% formalin for 6 months. For additional analyses involving antigen preparation or multi-omics of L3-liver, L3-lungs, or L4-intestine stages, wash the larvae thoroughly in PBS without fixing them. For ultrastructure-expansion microscopy (U-ExM) of Ascaris larval stages, refer to Basic Protocol 5.

• 30Transfer a small aliquot of the resuspended larvae suspension to a microscopy slide and observe under a microscope to quantify the larval stages.
• 31Repeat this step with all larvae suspensions.

Figure 7. Larval recovery from mouse tissue.

Modified Baermann apparatus for Ascaris larval recovery from chopped tissues, using a 100μm strainer and 50mL conical tubes.

BASIC PROTOCOL 4:

IN VITRO ASCARIS L3 LARVAE HATCHING AND PURIFICATION

Several studies have characterized the transcriptional profile and antigen repertoire of different stages of helminth parasites, including Ascaris suum and Ascaris lumbricoides (Gonzalez-Miguel et al., 2014; Phuphisut et al., 2022; T. Wang et al., 2013; Z. Wang et al., 2013), and have demonstrated their importance for the up- or down-regulation of host immunity, and their potential as vaccine and therapeutic targets. Thus, to investigate molecular patterns, antigenic diversity, and immunogenicity of the infective L3 larval stages, it is necessary to isolate them from the eggs. In this way, the next guideline is to standardize an in vitro Ascaris larval hatching protocol from fully embryonated eggs, based on an abrupt change of the pH of the egg solution, and further larval purification.

Materials

Ascaris fully embryonated eggs (Basic Protocol 1)

5% Sodium hypochlorite

Sterile DI water

Hank’s Balance Saline Solution (pH 2.0)

Phosphate-buffered saline – PBS (pH 7.4)

RPMI 1640 media (Gibco, cat.# 11875135) supplemented with 1% Penicillin-Streptomycin (Gibco, cat.# 15140122)

15 mL and 50 mL conical tubes

Serological pipette

CO₂ incubator at 37°C

Centrifuge

24-well culture plate

12-well culture plate

20 μm strainers (pluriSelect, cat.# 43-50020-03)

1 mL pipette

Incubator at 37°C

pH meter

Biological Safety Cabinet (BSC)

Protocol steps with step annotations

1. Inside the BSC, transfer approximately 10 mL of the fully embryonated egg suspension (See Basic Protocol 1) to a 50 mL tube.

2. Centrifuge at 1000 rcf for 10 minutes at 25°C.

3. Discard the supernatant using a serological pipette.

4. Resuspend the pellet of eggs in 25 mL of 5% sodium hypochlorite.

5. Incubate for 2 h at 37°C and 5% CO₂.

6. Perform 2 washes with sterile DI water (1000 rcf for 10 minutes at 25°C).

7. Resuspend the pellet in 15 mL of sterile Hank’s Balance Saline Solution (HBSS) pH 2.0.

8. Incubate for 30 minutes at 37°C and 5% CO₂.

9. Add 30 mL of PBS (pH 7.4) to the top of the egg suspension.

10. Centrifuge at 1000 rcf for 10 minutes at 25 °C.

11. Resuspend the pellet containing the fully embryonated eggs in 40 mL of RPMI media (pH 7.0–7.4).

12. Centrifuge at 1000 rcf for 10 minutes at 25 °C.

13. Resuspend the pellet with 3 mL of RPMI 1640 media supplemented with 1% antibiotic.

14. Transfer each 1mL of the eggs suspension in separate well of a 24-well plate.

15. Every 24 h for 10 days: collect one 20 μL aliquot per well and analyze the larval hatching ratio by light microscopy (non-embryonated eggs, eggs with larvae inside, and hatched larvae) (Figure 8).

    NOTE: Alternatively, to produce excretory/secretory ES antigens from infective L3 larval stages, while checking the hatching ratio from days 4 to 9 (once the ratio of hatched larvae is above 50%), collect only the media in the supernatant of the well and add new fresh media. After pooling all the collected media together containing all ES antigens produced by the larval stages, concentrate it using a 3 kDa centricon, then quantify their protein content using BCA, Lowry or Bradford assays, or characterize them using molecular or immunological assays, such as ELISA or Western Blott assays.

16. The hatched larvae is purified in a 12-well culture plate.

17. Insert a 20 μm strainer into one of the wells of the 12-well plate.

18. After at least 5–7 days of hatching, transfer the culture suspension containing the mix of eggs and hatched larvae into the strainer and fill up the well and strainer with RPMI media supplemented with 1% antibiotics (total volume of 10 mL/well)(See Figure 9).

19. Place the plate with the strainer in the incubator and incubate overnight, allowing all the larvae to migrate to the bottom of the well (since it’s a 20 μm strainer, only hatched larvae can pass through in a vertical position).

20. Check for the presence of purified larvae in the bottom of the well using an inverted microscope (See Figure 10).

21. Remove the media containing the eggs on the top of the strainer, carefully discard the strainer, and transfer all the volume inside the well containing the purified larvae to a 15 mL conical tube.

22. Centrifuge at 1500 rcf for 10 minutes at 4°C (low temperature helps larvae to pellet).

23. Remove all the supernatant with a 1 mL pipette.

24. Resuspend the pellet in 1 mL of PBS.

25. Count the larvae.

Figure 8: Ascaris eggs hatching ratio.

Three aliquots of the culture suspension were analyzed to calculate the larval hatching ratio under light microscopy. The graph represents the in vitro analysis of the frequency of non-embryonated eggs (grey), embryonated eggs (beige) and Ascaris hatched larvae (green), during a 9 day-period of culture.

Figure 9: Hatched larvae purification.

20μm strainers are used to purify hatched L3i larvae from the remaining eggs in culture. Only L3i stages are capable of passing through the 20μm filter during an overnight incubation at 37°C, 5%CO₂. strainer. Eggs will be retained.

Figure 10: Purified hatched L3i stages.

Left: Ascaris fully embryonated eggs before the hatching protocol. Scale – 20μm. Right: purified L3 hatched larval stages, after the utilization of the above protocol. Scale - 50μm.

SUPPORT PROTOCOL 1:

CRUDE ANTIGEN PREPARATION FROM ASCARIS PARASITE STAGES

The following protocol describes the preparation of crude extract soluble antigens from four stages of the life cycle of Ascaris spp., including infective L3i, L3-lungs, L4-intestinal larval stages and adult worms.

Materials

Lung-stage Ascaris larvae purified from BAL (Basic Protocol 3)

Intestine-stage Ascaris L4 larvae (Basic Protocol 3)

Ascaris infective L3 larval stages purified from eggs (Basic Protocol 4)

Phosphate-buffered saline (PBS)

Benchtop centrifuge

Polypropylene Tissue Grinder (Axygen^™, cat.#14-222-358)

Ultrasonic Probe Tip Sonicator (Qsonica, Q55, cat.#4422)

Bucket with ice

5 mL conical tube

1.5 mL microtube

Bicinchoninic Acid (BCA) Protein quantification kit (Thermo Scientific, Pierce^™, cat# A55864)

Protocol steps with annotations

1. Use either L3-infective larval stages recovered from eggs (Basic Protocol 4), and/or L3-lung stages recovered from airways in the BAL (Basic Protocol 3), and/or L4-intestinal stages recovered from the small intestine (Basic Protocol 3), and/or adult stages (Basic Protocol 1).

2. In a 5 mL conical tube, gather from 1mL of volume of the larval stages or use around 100 mg of adults, and macerate with a disposable tissue grinder in PBS.

3. Sonicate the homogenate of the parasite, using a probe tip sonicator at a frequency of 60Hz. Set a program of 10 cycles of 60 seconds, with 30 seconds of break between each cycle. Do the entire sonication process with the tube placed on ice.

4. Centrifuge at 3000 rcf for 15 min at 4°C.

5. Collect the supernatant and store it in a 1.5 mL conical tube.

6. Quantify the protein content in the crude antigen by Bicinchoninic Acid (BCA).

BASIC PROTOCOL 5:

ULTRASTRUCTURE-EXPANSION MICROSCOPY (U-ExM) OF ASCARIS SUUM LARVAL STAGES.

Light microscopy is a key tool in the study of fundamental cell biology. During their larval stages, however, the small size of Ascaris larvae can make observing individual cells or subcellular structures challenging using conventional light microscopy. Further, nematodes possess a cuticle that is largely impermanent to antibodies leading to difficulty in staining intracellular proteins (Duerr, 2013). One technique that is now widely applied to overcome the small size of cells or biological structures is ultrastructure-expansion microscopy (U-ExM) (Gambarotto et al., 2019; Hinterndorfer et al., 2022; Langner et al., 2024; Liffner & Absalon, 2024). U-ExM physically expands a biological sample while both preserving its anatomical an ultrastructure. Here we detail a protocol for U-ExM (Figure 11) of L3-lungs and L3-infective Ascaris larvae (isolated using Basic Protocols 3 and 4) that results in approximately 2.3-fold expansion of larvae (Figure 12). Many anatomical features can easily be observed in larvae prepared by U-ExM, including the nerve ring, excretory pore, intestinal microvilli, and esophago-intestinal valve (Figure 13 and 14). This protocol was derived from one describing expansion microscopy of Caenorhabditis elegans (Yu et al., 2020), with adaptations derived from a protocol on U-ExM of mosquito tissues (Liffner et al., 2024).

Figure 11. Workflow diagram for U-ExM of Ascaris larvae.

Note that the workflow alternates between read from left to right (steps 1. – 4. And 6. – 9.) and right to left (steps 5. And 10. to 11.) between rows. PFA: paraformaldehyde, conc: concentrate by centrifugation at 2000 rcf for 7 minutes, PG = propyl gallate, PDL = Poly-D-lysine. Snowflake symbols indicate points during the protocol where gels can be cryopreserved.

Figure 12. Validation of expansion factor by measuring nucleus diameter.

L3-infective and L3-lungs larvae were stained with dyes for nucleic acid (, SYTOX) and protein density (greyscale, NHS Ester) and imaged by Airyscan Microscopy either unexpanded or after U-ExM. To estimate larval expansion factor, the maximum nucleus diameter was measured across multiple nuclei in multiple larvae. Colour-matched datapoints represent nucleus measurements from the same larva. Small datapoints represent individual nucleus measurements while large datapoints represent the mean of each larva. Error bars = SD.

Figure 13: Anatomical features of L3-infective larvae prepared by U-ExM.

L3-infective larvae were prepared by U-ExM, stained with dyes for nucleic acid (, SYTOX) and protein density (greyscale, NHS Ester) and imaged by Airyscan Microscopy. Specific anatomical features are outlined in either or . Note that not all images of individual features come from the larva depicted in the whole image. Black scale bar = 50 μm, white scale bar = 5 μm.

Figure 14: Anatomical features of L3-lungs larvae prepared by U-ExM.

L3-lungs larvae were prepared by U-ExM, stained with dyes for nucleic acid (, SYTOX) and protein density (greyscale, NHS Ester) and imaged by Airyscan Microscopy. Specific anatomical features are outlined in either yellow or magenta or have arrows pointing to them. Note that not all images of individual features come from the larva depicted in the whole image. Yellow scale bar = 200 μm, Black scale bar = 20 μm, white scale bar = 10 μm.

Materials

Ascaris L3-infective or L3-lungs larvae from Basic Protocols 3 or 4

Deionised, nuclease-free water

Paraformaldehyde 4% (PFA) in PBS (Fisher, cat.# AAJ61899AK)

Poly-D-Lysine 0.1mg/mL solution (Gibco, cat.# A3890401)

Triton^™ X-100 (Tx100), 98% (ThermoFisher cat.# 327371000)

1.5 mL tube cap locks (Fisher cat.# NC9679153)

12 mm round coverslips (Fisher cat.# NC1129240)

35 mm Cellvis #1.5 glass bottomed dishes (Fisher cat.# NC0409658)

37 °C incubator

Dissecting forceps (Fisher cat.# NC9889584)

Dry heat block

Humid chamber

Orbital shaker

Parafilm (Sigma cat.# P7793)

Smart plastic razors (Sigma cat.# Z740503)

Thin paintbrush (#2 to #4 brush width)

Tube rotator

Vortex

Protocol steps with step annotations:

1. In a 1.5 mL tube, fix either L3-infective (prepared using Basic Protocol 4) or L3-lungs (prepared using Basic Protocol 3) larvae in 4% PFA in PBS for 1 hour at RT.

2. Concentrate larvae by centrifugation at 2000 rcf for 7 minutes. Remove fixative, resuspend larvae in 1 mL cuticle reduction buffer, and incubate overnight at 4 °C on a tube rotator.

3. Concentrate larvae by centrifugation. Wash twice with cuticle wash buffer and twice with wash buffer before resuspending larvae in 1 mL of anchoring solution. Anchor overnight at 37 °C.

4. Thaw TEMED and APS aliquots on ice for ~30 minutes. Prepare humidity chamber with one square of parafilm for each gel that will be made and store humidity chamber at −20 °C for ~15 minutes.

5. Remove humidity chamber form −20 °C, place on ice and place a poly-D-lysine coated round 12 mm coverslip on each square of parafilm.

6. Concentrate larvae by centrifugation, wash once in PBS and resuspend in ~ 15 μL of PBS for each coverslip to be prepared. Transfer 15 μL of larvae in PBS to each coverslip and allow to partially evaporate for ~ 5 minutes.

7. Add 5 μL of APS and 5 μL of TEMED to an aliquot of monomer solution and quickly vortex. Place 35 μL of activated monomer solution onto each square of parafilm (2 gels per aliquot). Using forceps, flip the coverslip onto the activated monomer solution (larvae side facing the gel).

8. Allow gel to partially polymerise on ice for 5 minutes, before transferring humidity chamber to 37 °C incubator for 30 minutes.

9. Place polymerised gels into the wells of a 6-well plate containing 2 mL denaturation buffer and place on a shaker for 15 minutes to detach the gel from the coverslip.

   NOTE: It is easiest to manipulate gels with the tip of a thin paint brush (similar to the kind specified in the materials)

10. Place each gel into a 1.5 mL tube filled with denaturation buffer. Place a safety cap on the tube and incubate in a heat block at 95 °C for 90 minutes.

11. Following denaturation, expand gels by transferring to a 10 mL dish filled with MilliQ water and place on a shaker for 30 minutes at RT. Perform two further water washes for 30 minutes.

    NOTE: Following the first expansion step, gels can be prepared for long-term cryopreservation by washing 3x in 50% glycerol and storage at −20 °C. To thaw gels, remove from −20 °C, wash for 30 minutes in MilliQ water and continue the protocol from Step 12.

12. Shrink gels back down by washing twice in PBS for 15 minutes. Full shrunken gels can be placed in a 6-well plate for staining or cut into smaller pieces and places in a 12 or 24-well plate.

13. Place gels in 1 mL blocking solution for 15 minutes at RT before adding primary antibodies diluted in blocking solution. Once in primary antibody solution, incubate overnight at RT on a shaker.

    NOTE: For full gels in 6-well plates, use 1 mL primary antibody solution. 500 μL for 12-well plates and 250 μL for 24-well plates.

14. After primary antibody incubation, wash gels three times in wash buffer for 10 minutes. Prepare secondary antibody solution (along with dyes like NHS Ester or DAPI) in PBS and add to gels for 2.5 hours at RT on a shaker in the dark.

15. Wash gels three times in wash buffer for 10 minutes and transfer back to 10 mL dishes with MilliQ water for expansion. Expand by washing three times in water for 30 minutes at RT.

    NOTE: If gels were cut earlier, the second expansion can be performed in a 6 or 12-well plate.

16. Place stained gel on a poly-D-lysine coated coverslip-bottomed 35 mm imaging dish and image.

17. If gels are not being imaged immediately, they can be placed in 0.2% propyl gallate and stored 4 °C for approximately 1 week.

Reagents and solutions

Cuticle reduction buffer [25 mM borate buffer (ThermoFisher cat.# J6902.AP)], 0.5% Tx100, 2% ß-mercaptoethanol [(ThermoFisher cat.# 125472500) in PBS)]

Cuticle wash buffer (25 mM borate buffer, 0.5% v/v Tx100 in PBS)

Sodium acrylate solution [(Sigma, cat.# 408220) 38% wt/wt in MilliQ water]

• WARNING: Use in fume hood.

TEMED solution [(ThermoFisher cat.# 17919) 10% in MilliQ water]

• WARNING: Use in fume hood.

• Aliquot and store at −20 °C

• Ammonium Persulfate solution (APS) [(ThermoFisher, cat.# 17874) 10% in MilliQ water]

• Aliquot and store at −20 °C

Monomer solution (sodium acrylate 19% wt/wt, acrylamide (Sigma, cat.# A4058) 10% v/v, BIS (Sigma, cat.# M1533) 0.1% v/v, in PBS)

• WARNING: Use in fume hood.

• Monomer solution MUST be prepared at least 24 hours in advance and can be stored at −20 °C for up to 2 weeks

• We typically make 900 μL of monomer solution as follows, and store as 10 × 90 μL aliquots (good for making 20 gels)

  • 500 μL 38 % wt/wt sodium acrylate solution
  • 250 μL acrylamide
  • 50 μL BIS
  • 100 μL 10x PBS

Denaturation buffer (200 mM SDS, 200 mM NaCl, 50 mM Tris, pH 9, in water)

• WARNING: Irritating to skin.

Anchoring solution [1.4 % v/v formaldehyde (Sigma, cat.# F8775), 2 % v/v acrylamide in PBS]

• WARNING: Prepare in fume hood.

Propyl gallate solution [0.2 % wt/v propyl gallate (ThermoFisher, cat.# 131581000) in MilliQ water]

Freezing solution [50% v/v glycerol (Fisher cat no: BP229-4) in MilliQ water]

Blocking solution (3% w/v bovine serum albumin in PBS)

Wash buffer [0.5% v/v TWEEN-20 (Sigma cat.# P1379) in PBS]

COMMENTARY

Several studies in the literature have shown that the transient migration of Ascaris larvae to mouse lungs leads to diffuse lung infiltrates and eosinophilic pneumonia (Weatherhead et al., 2020). These authors have demonstrated that during primary exposure to Ascaris, the migration of L3-larval stages from the pulmonary circulation to the lung parenchyma and subsequent penetration into the airways cause significant bleeding and mechanical damage to the lungs (Figure 15). The disruption of the epithelial barrier by the lung-stage migrating larvae triggers an inflammatory response resulting in a marked production of CXCL-1 driven neutrophils and increased levels of IL-6 (Gazzinelli-Guimaraes et al., 2013). L3 larval stages change in size and transcriptional program during their quest to the airways and elicit a dominant pulmonary type 2 immunity, characterized by elevated levels of IL-5, followed by the production of IL-4, IL-13, and CCL-11 (Eotaxin-1) (Figure 16). This cytokine/chemokine storm driven by Ascaris parasites in the lung tissue culminates in the differentiation of M2 macrophages and marked influx and activation of eosinophils, dependent on sub-epithelial fibroblasts and epithelial cells-mediated eotaxins (Gazzinelli-Guimaraes et al., 2019; Gazzinelli-Guimaraes et al., 2023).

Figure 15. Photograph of an Ascaris suum-infected mouse lung during the peak of pulmonary larval migration at day 8 of infection.

Observe the macroscopic hemorrhagic spots on the lung surface, resulting from mechanical damage caused by larval migration.

Figure 16. Cytokine profile of Ascaris-infected mice in different tissues.

Scatter plots show the sera levels (A), and tissue-specific cytokine [lung homogenate (B) and BAL fluid (C)], profile of naïve (blue) and Ascaris suum-infected BALB/c mice (red) at day 8 of infection. Each dot represents a single mouse, and the data is demonstrated as the median + SEM.

Long-term consequences of the immunopathogenesis driven by pulmonary larval migration include asthma and potentially chronic lung disease. Using the Ascaris mouse model, Weatherhead and collaborators (Weatherhead et al., 2018) demonstrated lung ascariasis resembles an extreme form of allergic airway disease with persistent airway hyperresponsiveness (AHR). In addition, this model has been used to unravel the mechanisms to explain how Ascaris infections may induce or exacerbate allergic inflammation by assessing the impact of the cross-reactivity between allergens- and Ascaris-antigens in the allergic effector response, as well as in the mediated resistance to helminths. Recently, investigating the interface between Ascaris infection and pulmonary allergic inflammation induced by house dust mite (HDM), we identified Ascaris-encoded tropomyosin and enolase as the two major HDM homologues based on high sequence and structural similarity (P. H. Gazzinelli-Guimaraes et al., 2021). We showed that the HDM-triggered IgE cross-reactive antibodies were functional as they mediated immediate hypersensitivity responses in skin testing. We also demonstrated that helminth tropomyosin was capable of inducing a severe type-2 associated pulmonary inflammation following the sensitization with the homologous house dust mite tropomyosin (also known as Der p 10).

These studies that have been primarily focused on understanding the pathobiology of larval ascariasis and the potential players associated with disease and the elicited protective immunity against Ascaris parasites have the potential to significantly contribute to the development of prophylaxis strategies and the discovery of new therapeutic targets capable of blocking or interrupting the pathogenesis of the disease before the establishment of chronic infection.

Critical Parameters:

1. The peak of Ascaris egg infectivity for experimental infections varies between 100–150 days in culture, however, the eggs become infective around day 35. After 200 days in culture, the egg’s infectivity reduces dramatically (Gazzinelli-Guimaraes et al., 2013).

2. Proficiency in animal handling is essential for gavage administration in mice. To minimize variability within the infected group, ensure to mix the egg suspension immediately before each syringe drawing, as the embryonated eggs settle rapidly. Prepare the egg suspension and perform the infection on one mouse at a time.

Troubleshooting:

Table 1 provides a detailed guidance, including problems, their possible causes, and recommended solutions to help you to address potential challenges/issues encountered while conducting some of the protocols related to egg embryonation culture, infection procedures, and ultrastructure-expansion microscopy (U-ExM) of Ascaris suum larval stages.

Table 1:

Troubleshooting guide

Problem	Possible cause	Solution
The volume of the flasks containing embryonating eggs gradually decreases in the incubator over time.	Prolonged incubation of Ascaris eggs for embryonation at 26°C.	Regularly supplement the culture flask with additional PBS or sulfuric acid solution on a weekly basis to prolong its viability.
Reduced larval burden or increased variability in larval burden within the tissues.	During the infection, the embryonated eggs tend to settle rapidly in the tubes.	Only gather the necessary volume of eggs in the syringe (200 μL) for each individual mouse. Avoid collecting 1 mL of eggs in the syringe for a group of 5 mice.
Gel is not fully expanding.	Old/faulty sodium acrylate.	There is significant batch-to-batch supplier variation in SA. Check SA isn’t yellow when put into solution. Replace approx. every 6 months.
Gel is not fully expanding.	Old monomer solution.	Monomer solution will start to produce inconsistently sized gels after approx. 2 weeks.
Larvae not reaching ~2.5-fold expansion.	Incomplete cuticle reduction or denaturation.	Ensure overnight reduction in cuticle reduction buffer. Ensure direct contact between 1.5 mL tube and heat source during denaturation.
Larvae too deep into the gel to image.	Imaging the ‘wrong’ side of the gel.	The side of the gel in contact with the parafilm will have a ‘grainy’ pattern. The larvae will be closer to the other side of the gel, which was in contact with the coverslip.
Gel is drifting when you image.	Not poly-D-lysine coated.	Ensure imaging dishes have been coated with poly-D-lysine. Poly-L-lysine is not sufficient for this.

Understanding Results:

1. Lewis and collaborators (Lewis et al., 2006) have demonstrated a strong positive correlation between the size of the inoculum and the mean worm recovery from the lungs in both Ascaris-infected CBA/Ca and C57BL/6j mice. In addition, we present original results to demonstrate that the immunopathogenesis of larval ascariasis in the lung tissue homogenate follows a dose-response manner according to the intensity of infection (Figure 17).

2. Studies in the literature have demonstrated that primary infection in naïve mice and secondary infection by repeated inoculations (a way to represent the multiple exposures in the endemic areas) provide completely different outcomes for parasite burden in the tissue, and for the elicited immunopathogenesis. Nogueira et al. (Nogueira et al., 2016) demonstrated that using the protocol highlighted in Figure 18, multiple exposures to Ascaris parasites in mice elicited an intense eosinophil-dominated pulmonary inflammation associated with a polarized mixed Th2/Th17 immune response which was associated with a 90% reduction in the parasite burden when compared with a primary infection in naïve mice.

3. It was demonstrated (Deslyper et al., 2016; Deslyper, Holland, et al., 2019) that there was a connection between the genetic background of the mice and their inherent susceptibility (or lack thereof) to Ascaris infection. Lack of susceptibility (i.e., resistance) was associated with oxidative phosphorylation pathways, reactive oxygen species production and expression of components of the complement system in the liver.

Figure 17: The cytokine profile of Ascaris-infected lungs correlates with larval intensity in the tissue.

BALB/c mice were infected either with 500, 2500 or 10000 eggs, or non-infected. (A) Lung parasite burden of mice infected with different doses of Ascaris suum eggs at 8 days post-infection. (B) Lung tissue homogenate of mice infected with different doses was used to measure cytokines/chemokines by Luminex assay. The heatmap showcases the levels of specific key cytokines in the context of varied egg doses, highlighting the dose-dependent immune responses. P values and Spearman r values for each cytokine/chemokine are indicated in the graph.

Figure 18: Experimental model for re-infection with Ascaris parasites.

In the re-infection model, mice receive multiple exposures by three intragastric administrations of 2,500 fully embryonated eggs spaced by 14 days apart at days −28, −14 and 0. Following the final infection, mice are euthanized, and tissue collection time points are focused on the sites to which larvae migrate (liver at 4, lugs at 8 and intestine at 12 days post-infection). This figure was made using Biorender.com.

Time Considerations:

The time required for isolating eggs from the parasite female’s uterus will vary depending on the number of adult worms present. Anticipate the entire process to take approximately 2 to 4 hours.

When euthanizing mice and collecting organs, it’s best to begin early in the day. The duration of euthanasia and collection can vary based on the number of participants and their expertise. The intestine takes the longest to collect, so we suggest starting with bronchoalveolar lavage (BAL), then proceeding to the liver, lungs, and finally the intestine. Collecting all organs from one animal typically takes between 10 to 40 minutes.

For larval burden assessment, allocate additional time since the counting is manually using a light microscope. Counting larvae in the BAL is quicker owing to cleaner samples. However, expect longer counting times for larvae in other organs due to the presence of debris. Plan for approximately 10 to 30 minutes for counting larvae in each sample.

DATA AVAILABILITY STATEMENT:

The data that support the findings of this study are available from the corresponding author upon request.