Symbiosis is a fascinating and diverse phenomenon. The study of symbiosis is important to understanding ecology, as it helps us understand relationships between organisms and provides insight into co-evolution, mutualism, adaptation, and survival. Ecological studies are challenging to implement in K-12 classrooms because they often require multiple organisms (often very different in size) and complex environments that are difficult to replicate accurately (e.g., soil composition, temperature, pH, and humidity). These factors can make it difficult to study quantitative changes in ecosystems. We developed an inexpensive, quantitative experiment for classrooms that can be used to explore important aspects of microbial symbiosis, pathogenesis, and ecology, and that helps support more investigations in this area of education. The experiment is low-cost, designed for K-12 teachers and students, uses common materials, and teaches students about the exciting relationships among bacteria, worms, and insects.

Introduction

Symbiosis—the interaction between different species—is found in every ecosystem studied to date (Saffo, 2014). To fully understand an ecosystem, one must look at all of the interacting partners to see how the actions of one species affect another. Most well-known examples of these interactions—e.g., sea anemones and clown fish (Fautin, 1991; Miller, 2016), or oxpecker and rhinoceros (Mengesha, 1978)—are difficult to bring into a classroom. Microbes, on the other hand, are usually inexpensive and easy to acquire and grow. However, the small size of microbes (typically 1–50 microns) can make them difficult to study, especially when microscopes are not available. Nematodes (roundworms) are large enough to see with the naked eye (~1 mm) and are inexpensive and easy to grow. They are also involved in several interesting interactions with other organisms, making them ideal for studying symbiosis.

Nematodes have adapted to almost every ecological niche on the planet, and are so abundant that four out of five animals on the planet are nematodes (Chen et al., 2004). Nematodes form mutualistic (mutually beneficial) relationships with bacteria to acquire nutrients. Although numerous species are animal or plant parasites, many nematodes and their bacterial symbionts are useful for insect pest control in agriculture, making them excellent model organisms for academic studies of symbiosis (Dillman & Sternberg, 2012; Ehlers, 2001). For example, the soil-dwelling, entomopathogenic (insect-eating) nematode Steinernema feltiae forms a symbiotic relationship with the bacterium Xenorhabdus bovienii, which helps it kill and digest its insect prey (Hirao & Ehlers, 2009). S. feltiae navigates through soil toward its prey by responding to concentration gradients of chemical signals, or chemoeffectors, released by its insect prey in a process called chemotaxis (chemo as in chemical, taxis as in movement) (Hui & Webster, 2000).

It is easiest to study nematodes when their movement is confined. We use small microfluidic channels to accomplish this. These channels are shallow enough that their motion is essentially limited to two dimensions, making them easier to observe (San-Miguel & Lu, 2013). Microfluidics experiments use very small volumes of fluids (down to 10−18 liters) in channels that have a width or height that is on the scale of micrometers (1 micrometer is one millionth of a meter) (Whitesides, 2006). Scientists and engineers use microfluidic technologies to reduce the size, cost, and materials needed for experiments ranging from chemical synthesis to DNA sequencing (Abate et al., 2013; Elvira et al., 2013; Feng et al., 2015). The constraint imposed by microfluidic channels creates a phenomenon referred to as laminar flow (see goo.gl/BxIHNf for a demonstration of this property). Fluids exhibiting laminar flow move in “sheets,” which differs from the turbulent flow that is often observed in mixing fluids (Figure 1) (Falkovich, 2011). Laminar flow can be used to manipulate fluids very precisely (Whitesides, 2006).

Figure 1.
A depiction of laminar flow and turbulent flow. (A) Laminar flow causes fluids to move in sheets such that the fluids only mix by diffusion. (B) Turbulent flow causes fluids to readily mix and is the state we commonly observe when watching liquids. A convenient way to adjust between laminar and turbulent flow is by adjusting the diameter of a channel, d.
Figure 1.
A depiction of laminar flow and turbulent flow. (A) Laminar flow causes fluids to move in sheets such that the fluids only mix by diffusion. (B) Turbulent flow causes fluids to readily mix and is the state we commonly observe when watching liquids. A convenient way to adjust between laminar and turbulent flow is by adjusting the diameter of a channel, d.

Microfluidics is a powerful method to study chemotaxis, but traditional methods for creating microfluidic channels are often very expensive and require resources that are not commonly available (Xia & Whitesides, 1998). We use a technique that requires only inexpensive office supplies (double-sided adhesive and transparency sheets) and a craft cutter (Clawson et al., 2018). Using these microfluidic channels, we demonstrate an easy way to measure the chemotaxis of nematodes in response to various chemicals. Using this approach, students can ask whether nematodes swim toward or away from different chemicals—including those released by insects. This gives them the tools to answer fundamental ecological questions about nematode behavior—in this case, how the nematode S. feltiae identifies and hunts insect prey.

The Nematode Lifecycle

Nematodes can infect a wide range of hosts, including insects, plants, and mammals (Lee, 2002). Entomopathogenic nematodes, such as S. feltiae, can infect and consume a wide range of insects (Lacey & Georgis, 2012). S. feltiae has a mutualistic relationship with the bacterium Xenorhabdus bovienii, which they carry in specialized pockets in their intestines (Hirao & Ehlers, 2009; Kim et al., 2012). S. feltiae senses chemical compounds produced and released by the insect prey and uses this information to track prey (Figure 2) (Hui & Webster, 2000). After catching its prey, the nematode burrows into natural openings in the insect such as its mouth, anus, or spiracles (breathing openings) (Dowds & Peters, 2002). Once inside, nematodes release their symbiotic bacteria, which multiply and produce toxins and other compounds that kill and help digest the insect, which is then eaten by the nematode and the bacteria to provide energy for survival and reproduction. After the insect nutrients are depleted, the nematodes re-associate with the bacteria, exit the insect cadaver, and traverse the soil in search of their next insect prey (Figure 3) (Dowds & Peters, 2002).

Figure 2.
A cartoon depicting nematodes responding to chemical signals released by prey insects. Each prey insect produces and secretes multiple organic compounds that nematodes chemotax toward and use to hunt the insect. Wavy lines indicate production of volatile compounds. Examples of compounds shown are (from top left, clockwise): α-pinene, C10H16 (waxworm), dimethylsulfone, C2H6O2S (house crickets), furan, C4H8O (earwig), acetone, and C3H6O (pillbug). Note that although we depict only one compound for simplicity, insects release many organic compounds.
Figure 2.
A cartoon depicting nematodes responding to chemical signals released by prey insects. Each prey insect produces and secretes multiple organic compounds that nematodes chemotax toward and use to hunt the insect. Wavy lines indicate production of volatile compounds. Examples of compounds shown are (from top left, clockwise): α-pinene, C10H16 (waxworm), dimethylsulfone, C2H6O2S (house crickets), furan, C4H8O (earwig), acetone, and C3H6O (pillbug). Note that although we depict only one compound for simplicity, insects release many organic compounds.
Figure 3.
A cartoon depicting the life cycle of nematodes. Nematodes traverse soil in search of insect prey, infect the insect, and subsequently release their symbiotic bacteria to kill the insect. The nematodes and bacteria feed on the dead insect until the nutrients are depleted, at which point they re-associate and exit the insect cadaver in search of new prey.
Figure 3.
A cartoon depicting the life cycle of nematodes. Nematodes traverse soil in search of insect prey, infect the insect, and subsequently release their symbiotic bacteria to kill the insect. The nematodes and bacteria feed on the dead insect until the nutrients are depleted, at which point they re-associate and exit the insect cadaver in search of new prey.

Method Development

Microscopic organisms move differently from larger ones (Purcell, 1977). Steinernema nematodes move by pushing their body against solid objects, which makes it difficult to observe nematodes swimming in water (Park et al., 2016). We filled channels with an agar solution (agar powder is available online and at brick-and-mortar stores), which produces a transparent gel in which the worms can swim (Hida et al., 2015). To form the gel, the agar solution must be boiled, then allowed to cool. We found the nematodes could survive brief exposure to the high temperature of the solution of melted agar, and after the gel set the nematodes could easily swim through the gel. The agar solution cools and solidifies rapidly, so filling the channels may require a level of coordination that is too challenging for elementary school students.

We designed a microfluidic system consisting of two straight inlet channels (with inlet holes) intersecting with a single central channel and ending in an outlet hole; the channel system looked like the letter Y. We used three layers of material to form the channels (Figure 4): (1) a layer of double-sided adhesive tape with the channels cut into it; (2) a top layer of transparency film forming the channel ceiling with the shape and dimensions of the adhesive tape (this layer contained the inlet and outlet holes); and (3) a bottom layer of transparency film forming the channel floor with the shape of the adhesive tape but slightly longer to provide a handle. We used a craft cutter to cut the channels out of the double-sided adhesive and to cut the top and bottom layers out of transparency sheets. We assembled the system by pressing a transparency film layer on each side of the adhesive, then hot-gluing a small piece of Tygon tubing to one inlet (Figure 4). A microscope slide can be used instead of transparency film for the bottom layer of the channel if it makes visualization easier. For a very detailed description of the process for making the microfluidic system, see our recently submitted paper in The Journal of Microbiology and Biology Education (Clawson et al.). A step-by-step video can be found at https://youtu.be/BDFWlELvzJo.

Figure 4.
The structure of the microfluidic channels and their assembly. (A) The components of a single channel. (B) Carefully aligning the adhesive with the smaller piece of transparency. (C) After attaching the second transparency, the plastic tubing is hot-glued onto only one inlet. (D) The finished channel.
Figure 4.
The structure of the microfluidic channels and their assembly. (A) The components of a single channel. (B) Carefully aligning the adhesive with the smaller piece of transparency. (C) After attaching the second transparency, the plastic tubing is hot-glued onto only one inlet. (D) The finished channel.

When using the channels, we treated the tubing as a reservoir for the chemoeffector, to ensure the channels would not become filled with air. We filled the tubing reservoir with our chemoeffector of interest while preparing for subsequent steps. (Note: If the tubing prevents the channel from fitting onto a microscope, the tubing can be removed after filling the channels by pulling it off or using scissors.) After soaking the nematode-containing sponge in water, we mixed the nematode solution with the warm agar solution, pipetted it on the other inlet hole (without tubing), and drew the two solutions through the channel using a syringe positioned at the outlet (Figure 5). To create a better seal, we put a small piece of tubing on the tip of the syringe. After a few minutes, the channel was moved to the microscope to visualize nematodes.

Figure 5.
How the experiments are performed. (A) An image depicting all of the materials required for this experiment. (B) The tubing at one inlet is filled with chemoeffector. (C) The agar is loaded into the other inlet. (D) The fluids are drawn into the microfluidic channels using a tubing-tipped syringe. (E) An image of the laminar flow profile.
Figure 5.
How the experiments are performed. (A) An image depicting all of the materials required for this experiment. (B) The tubing at one inlet is filled with chemoeffector. (C) The agar is loaded into the other inlet. (D) The fluids are drawn into the microfluidic channels using a tubing-tipped syringe. (E) An image of the laminar flow profile.

Our microfluidic chemotaxis system differs from those reported previously in a seemingly small but important way: we do not load channels entirely with a solution of agar. Instead, we dissolve chemoeffectors in water (rather than in an agar solution) and use laminar flow to fill one half of the main channel with chemoeffector and one half with agar and nematodes. After the agar sets, half of the channel contains the agar gel, and the other half of the channel contains the chemoeffector solution. There are two primary advantages to this approach. First, the channels are much easier for students to load when agar is added to only one of the two inlets because it cools and solidifies rapidly. We found this task to be difficult, so younger students would certainly have trouble correctly loading the channels. Our second reason for loading chemoeffectors in water is that the nematodes swim well in the agar gel but not in the liquid, so chemotaxing nematodes are essentially trapped if they move to the chemoeffector side. This makes counting the worms easier. We have not observed any nematodes swimming into the chemoeffector area (liquid side) when a repellant is present, so we believe false positives are unlikely.

To test our system, we compared the nematodes’ responses to an attractant (waxworm extract), a repellant (acetic acid; i.e., white vinegar), and water (a control). In each experiment, we observed 4–12 nematodes in the main channel. (Note: The amount of water used to soak the nematode sponge can be adjusted to increase or decrease the number of worms suspended in liquid in each channel.) To quantify the nematode response to each chemical, we calculated the chemotaxis index (CI) using the following equation:

 
CI= # nematodes on chemoeffector sidetotal # of nematodes in channel

According to this equation, a perfect repellant will create a CI of 0, and a perfect attractant will yield a CI of 1. After performing the experiment in triplicate, we found that the waxworm extract yielded a CI of ~0.71, the acetic acid yielded a CI of ~0.03, and the water yielded a CI of ~0.14. (See Table 1 for individual experimental values, or Figure 6 for a photo of the channels and quantification of the nematode response). (Note that ideally, as a control, water would yield a CI of 0.5). We found that adding food coloring to the chemoeffector solution greatly helped us visualize the boundary between the agar and the chemoeffector, and made it easier for us to see that both solutions were loaded into the channel. We also observed that the laminar flow profile in channels containing both the warm agar solution and the waxworm extract was not always evenly distributed, most likely due to the evaporation of the isopropanol in the waxworm extract. An occasional wavy boundary formed between the two solutions, but will not affect the outcome of the experiment.

Table 1.
Nematode chemotaxis results using the microfluidic channels.
Number of nematodes in agar sideNumber of nematodes in chemoeffector sideChemotaxis Index
Waxworm extract 0.25 
0.89 
1.00 
Acetic acid, 5%
(white vinegar) 
10 
10 0.09 
Water 0.18 
10 0.23 
Number of nematodes in agar sideNumber of nematodes in chemoeffector sideChemotaxis Index
Waxworm extract 0.25 
0.89 
1.00 
Acetic acid, 5%
(white vinegar) 
10 
10 0.09 
Water 0.18 
10 0.23 
Figure 6.
(A) Five nematodes in a channel loaded with attractant (waxworm extract); the image was taken ~5 minutes after loading the chemoattractant using an LG® G3 smartphone and a platform microscope outfitted with the lens from a laser pointer (Yoshino, 2015). Arrows indicate live nematodes. (B) Chemotaxis indices calculated from three separate experiments.
Figure 6.
(A) Five nematodes in a channel loaded with attractant (waxworm extract); the image was taken ~5 minutes after loading the chemoattractant using an LG® G3 smartphone and a platform microscope outfitted with the lens from a laser pointer (Yoshino, 2015). Arrows indicate live nematodes. (B) Chemotaxis indices calculated from three separate experiments.

Materials

  • craft cutter (e.g., Cricut Explore One, $184.51; Silhouette Portrait, $149.99; or comparable craft cutter that allows uploading of designs)

  • Steinernema feltiae nematodes (e.g., Nemaglobe Fungus Gnat Control Nematodes, available from Amazon.com)

  • fresh waxworms (available at bait and pet stores or Amazon.com; can be frozen to maintain freshness)

  • mortar and pestle (available at most cooking stores or Amazon.com)

  • 70% isopropanol (available at drug stores and grocery stores)

  • 5% acetic acid (white vinegar)

  • agar powder (available on Amazon.com or at ethnic food stores)

  • transparency film (available at office supply stores) (Write-on sheets are less expensive, but laser-compatible sheets are necessary for lined transparency guides.)

  • double-sided adhesive (e.g., Elizabeth Craft Designs Clear Double-Sided Adhesive, 8.5 by 11 inch, available on Amazon.com)

  • 3-mm hole punch (e.g., Cmxsevenday No.97C3 Metal Handheld 1-Hole Metal Punch, 1/8″ Hole Size, available on Amazon.com) (Note that most hole punches are 6 mm and thus are too large.)

  • plastic tubing (3/16″ inside diameter) (e.g., Tygon B-44-3 PVC Beverage Tubing)

  • hot glue gun (available at craft stores)

  • 1 mL syringes, Luer slip tip (e.g., Easy Glide 1cc Luer Slip TB Syringe) (Larger syringes will work, as well as Luer-Lok tips, but smaller syringes will be easier to work with.)

  • food coloring (optional, but highly recommended)

  • transfer pipettes with gradations (200 μL-1000 μL range) (e.g., Karter Scientific Plastic Transfer Pipettes 1ml, Graduated), or variable volume micropipette(s)

  • water bath at 50–55°C (122–130°F) (A beaker of water on a hot plate will work.)

Methods

  1. Assemble the microfluidic channels as described in Clawson et al. (2018) and shown at https://youtu.be/BDFWlELvzJo. Cut the channels out of the double-sided adhesive, and cut the top and bottom out of the transparency sheet. Remove the paper backing from the double-sided adhesive and attach to the smaller transparency. Ensure holes are cut out of the transparency either by the craft cutter or manually using the 3-mm hole punch. Remove the paper backing from the other side of the adhesive and adhere to the larger transparency (Figure 4). Push all layers together from end to end to avoid creases.

  2. To create the chemoeffector reservoir and syringe adapter, cut two 0.5-0.75 cm pieces of tubing (we use Tygon PVC tubing, SAE, 3/16″; inside diameter, ¼″ outside diameter, 1/32″ wall). Hot glue one piece of tubing to one inlet to create a reservoir to hold the chemoeffector (Figure 4). Attach the other piece of tubing to the tip of the syringe to make pulling the solutions into the channel easier. Use caution with hot glue to avoid burns. Avoid sealing the access holes to the microfluidic channel.

  3. Prepare the chemoeffectors as outlined below:

    • Attractant: Waxworm extract. Use the mortar and pestle to grind 5 frozen waxworms (freeze waxworms while they are fresh). Suspend the waxworm mash in 3 mL of 70% isopropanol. Prepare a twenty-fold dilution (e.g., 1 mL mash + 19 mL water) from this solution using water. Add food coloring to visualize the liquid. (Note: If sacrificing insects is not acceptable for the experiment, chemically pure chemoeffectors can be purchased from scientific supply companies.)

    • Repellant: Acetic acid (vinegar). Make a solution of 5% acetic acid in water (or use white vinegar, which is ~5% acetic acid in water), and add a second color of food coloring to visualize the liquid.

    • Control: Water. Add a third color of food coloring to visualize the liquid.

    Adding different colors to the solutions will make it easier to see the laminar flow profile and clarify that the channel is working as expected.

  4. Suspend the nematode sponge in water. We resuspended one sponge of nematodes in ~100 mL water, or half of a sponge in ~50 mL water.

  5. Melt a 2% agar solution and keep warm in a water bath (approximately 55°C). For ease of viewing, add a fourth color of food coloring.

  6. For each group of students, prepare three microfluidic channels—one each for attractant, repellant, and control—as outlined above and in Clawson et al. (2018). Provide each group of students with assembled microfluidic channels (or have them assemble their own), along with the 1-mL syringe connected to the extra piece of tubing and some sort of pipet.

  7. One student loads a chemoeffector solution into the inlet tubing (as in Figure 5b). Meanwhile, a second student mixes the nematode suspension with the 2% agar solution in a 3:1 ratio, so that the final agar concentration is 0.5% (e.g., 0.5 mL of the nematode suspension with 1.5 mL of the 2% agar solution). Immediately after mixing, the second student pipets the nematode-agar mixture onto the other inlet hole (as in Figure 5c). As soon as the nematode solution covers the inlet hole, the first student quickly and gently draws the two solutions into the channel using the tubing-tipped syringe placed at the outlet hole (as in Figure 5d). Note that the agar solution cools quickly, so this step needs to be performed quickly.

  8. Repeat step 7 for the other two microfluidic channels and conditions.

  9. Image the channels using a microscope (Yoshino, 2015). Record the number of nematodes in each side of the channel over time (e.g., at 5, 10, 15 minutes). To quantify the nematode response to the chemoeffector, use the following equation to calculate the chemotaxis index (CI):

     
    CI=#nematodes on chemoeffector sidetotal # of nematodes in device

    A chemotaxis index of 1 indicates that the chemical was a strong chemoattractant, and a value of 0 indicates a strong chemorepellant.

Note: The nematodes can be disposed of by rinsing them down a sink or releasing them into soil outside. The used channels can be flushed with hot water for reuse or discarded.

In the Field

The experiment was completed by 76 middle school students lacking formal biology training. The students experienced this investigation in a field trip setting at a science center near our lab. The teachers selected this workshop for their students from a number of other field trip options. To gain insight into the activity's impact on the students’ enjoyment and engagement, we asked them to complete a short survey after the workshop. The survey was based on the learning activation surveys developed by Activation Lab (activationlab.org). The frame of reference for this survey was on looking at evaluative measures that would indicate how the activity impacted participants. The surveys were anonymous and optional. No identifying information was collected from the students, and all responses were aggregated. Based on these attributes, Institutional Review Board staff deemed these evaluative measures to be exempt from IRB process.

We found this activity to be a rich platform for collaboration and a reflection of the practices we do as scientists. For brevity, we aggregated responses from several questions addressing a similar concept (collaboration with others) into a single measure (Figure 7a). When asked in anonymous surveys what they enjoyed most about the activity, more than half the students said “everything” or “we learned by doing things instead of listening the whole time,” while the rest “enjoyed being able to see the ways the nematodes reacted.” We observed students talking among themselves about what they were doing and working together to do the activity successfully, in much the same way that academic scientists collaborate in the research lab. Additionally, over 90 percent of the students reported that they asked questions, tried out new ideas, and discussed the experiment with their mentors and peers to aid their comprehension (Figure 7a). These are all hallmarks of a collaborative process that the activity's framework provided.

Figure 7.
Student responses to survey questions regarding the activity. (A) Student responses to questions concerning collaboration and exploration (N = 76). Students reported that they were actively checking their own understanding of the experiment (N = 22). (B) Students were actively engaged in the experiment. The majority of students participated in the activity because they wanted to, not because they were instructed to (N = 76). Most of the students felt they learned something about science (N = 22).
Figure 7.
Student responses to survey questions regarding the activity. (A) Student responses to questions concerning collaboration and exploration (N = 76). Students reported that they were actively checking their own understanding of the experiment (N = 22). (B) Students were actively engaged in the experiment. The majority of students participated in the activity because they wanted to, not because they were instructed to (N = 76). Most of the students felt they learned something about science (N = 22).

The activity provided opportunities for high levels of engagement. Of all student responses, 80–90 percent indicated that they were actively engaged and self-reported that they had learned something about science (Figure 7b). This activity can span engagement across age bands. We found this activity to be appropriate for students as young as 6th grade and can be adapted for students in high school and college by exploring the key concepts more deeply.

The Role of Mutualistic Bacteria in Nematode Predation

Entomopathogenic nematodes form relationships with bacteria that enable nematodes to infect and digest their insect prey. S. feltiae nematodes work with X. bovienii bacteria to obtain nutrients from prey insects. To learn more about the mutualistic relationship between X. bovienii and S. feltiae, students can create and study the behavior of aposymbiotic nematodes (ASN)—worms that lack bacterial symbionts. For a detailed protocol on making ASNs, see McMullen and Stock (2014). To confirm that the ASNs do not contain Xenorhabdus, students can grind up nematodes and inoculate them onto Luria-Bertani (LB) media plates as outlined by Sanders (2012) and observe bacterial growth (Sicard et al., 2006). After obtaining ASNs, students can compare the ability of aposymbiotic worms to infect and kill prey insects with the ability of bacteria-colonized nematodes (as outlined in McMullen & Stock, 2014).

Conclusions

Using simple, inexpensive, easy-to-make microfluidic channels, we demonstrated how students can identify and study chemicals that alter the behavior of predatory nematodes, such as the chemicals that nematodes use in nature to track their prey and avoid danger. An important aspect of this activity is the engagement of students in the design of authentic scientific investigations. We often think of ecology on the scale of large animals and ecosystems; however, this activity reinforces the importance of ecosystems of small animals and bacteria, and enables teachers to investigate a complex and important ecological web using tools that are widely accessible and quantitative.

This activity can be expanded to fit within existing biology and environmental science curricula, as different aspects of symbiosis are reflected in this experiment: e.g., the mutualistic relationship between the worms and their symbiotic bacteria, as well as the pathogenic relationship between the worms and their insect prey. These concepts could fit in a curriculum studying bacteria, pathogens, chemical ecology, soil science, and insects. This activity can be modified to use different chemicals, temperatures, microfluidic channel shapes/designs, or species of nematode, and enables students to make predictions about the response of nematodes to those conditions. We have provided a worksheet for students to fill out as they perform the activity (Supplemental Material 1), as well as a worksheet (Supplemental Material 2) and key with provided data for practice analyzing data from the activity (Supplemental Material 3).

Microfluidics makes possible the investigations described here because it imposes dimensions on liquids that bring out the unique laminar behavior of fluids. Traditional methods for making microfluidic channels are tricky to incorporate into school activities because they require materials and facilities that are expensive and difficult to access. Our method uses inexpensive office materials, a craft cutter to pattern the double-sided adhesive and transparency sheets, and simple methods for assembly and introducing fluids. The ease of this method enables students to create and test new channel designs, and makes it possible to incorporate perspectives on engineering into lessons.

Educators will be able to incorporate the Educators Evaluating the Quality of Instructional Products (EQuIP) Rubric to measure the alignment to the Next Generation Science Standards (NGSS) to help determine which elements from these investigations connect to the science and engineering practices (SEP), disciplinary core ideas (DCI), and/or crosscutting concepts (CCC) of NGSS. The embedded tasks expected of students in these activities may demonstrate their proficiency of one or more performance expectations. For example, students develop and use SEPs, DCIs, and CCCs that fit within the EQuIP three-dimensional framework. Teachers will need to identify the key CCCs relevant to their instructional framework so that students can translate specific information to general principles.

Looking deeper into the EQuIP (v3.0) rubric, there are intentional opportunities for students to explain phenomena and design solutions that are integrated into life science DCIs (MS-LS2-1,2,4 & HS-LS2-8) and SEPs (MS/HS-ETS1). In particular, the evidence statements reflected in NGSS HS-LS2-8 connect very well to the potential for students to develop a causal explanation on the group behavior dynamics of chemotaxing nematodes in microfluidic channels (NGSS, 2015). Students will use science and engineering design practices in this activity and other potential investigations of ecosystems and inter-organism interactions.

We acknowledge funding from the Dreyfus Foundation (SG-10-032), National Science Foundation (DMR-1121288, AISL 1241429, and pre-doctoral fellowship DGE-1256259 to J.F.N.), Wisconsin Alumni Research Foundaton (MSN193090), and Madison Community Foundation (5947) that made this research possible. We are grateful to Heidi Goodrich-Blair and Mengyi Cao for introducing us to this fascinating area of biology.

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