# Appendix A: Water Bucket Protocol (WBP) #### The Water Bucket Protocol (WBP) Our team confesses a secret delight in such a simple name for such a complex problem as standardizing frontier-science eDNA analysis in remote field sites operated by Indigenous youth. The source of the idea is truly bi-cultural and multidisciplinary, as described in [Appendix B](/appendices/appendix-b-bicultural-design-and-co-authorship.md). NOTE: This protocol is in active development; what is written on this page will change and update in the proximal year to ongoing feedback from pilot testers (see Bricolage) #### **Purpose of the WBC** **The water bucket protocol standardizes eDNA data collection.** By using filtered water, a standardized time period, and plot centroids, it reduces noise and enhances signal in eDNA data collection. To recap the [Biodiversity calculation](/biodiversity/biodiversity-calculation.md) section, this offers a consistent approach for comparing sites over time to assess changes in ecosystem integrity. **The aim is to reduce costs and improve accuracy when using eDNA as a metric**. Results will need to be statistically normalized to a per-project standardized curve for integrity uplift over time. The WBC simply makes that easier by reducing the available data to a more meaningful set, reducing he sample size necessary to meet the output requirements for Interoperable Biodiversity Units (IBU) and qualify for recurring revenue from uplift biodiversity credits (roughly $300/unit, see [Biodiversity unit](/biodiversity/biodiversity-unit.md)). #### Limitations of the WBC There are serious practical constraints to field testing eDNA. The absence of established eDNA reference levels across ecosystems remains the primary challenge for using eDNA in biodiversity uplift assessments via IBUs, compounded by the logistical difficulties of exporting samples and the costs of analysis to obtain eDNA metrics. One possible approach is to establish a baseline sampling of eDNA in untouched or old-growth forests, representing conservation in Fig. 4C. Simple traditional biodiversity metrics, such as species richness, are well-suited for community-based monitoring but have low statistical power and thus require substantial sampling effort to reliably detect ecological changes [(Lamb et al. 2009)](https://doi.org/10.1016/j.ecolind.2008.06.001). Random error in eDNA samples may also be high and affect data integrity [(Lahoz-Monfort et al. 2016)](https://doi.org/10.1111/1755-0998.12486). Cost-effective monitoring thus requires statistically robust sampling, ideally with regional normalized curves — though these may be more expensive to develop in more biodiverse zones. That said, we believe in progressing limits where we find them, as practically as possible. The protocol below is intended to reduce sampling noise in a cost-effective way. #### **Sampling location** While our IBU calculations are designed for one-hectare areas, sampling is performed at the centroid of a plot (averaging five hectares) to optimize for the conditions of the agroforestry system itself rather than the biodiversity that tends to accumulate at the edges of farmed areas. Plot size should be used as a covariate in later statistical modeling (see[ Biodiversity calculations](/biodiversity/biodiversity-calculation.md)). #### **Materials** * Four locally-sourced buckets, sterilized * 10% bleach solution (for sterilization); clean water (for rinsing) * \~4 L of filtered water for filling * A sterilized, twice-rinsed 1 L sample bottle * Gloves and face masks * SimplexDNA filtration kit ([simplexdna.com](https://www.simplexdna.com/)) * KoboToolbox or equivalent for field data capture Bucket coloring should follow a consistent scheme: light colors to attract pollinators, dark or terrain colors to attract wood-dwelling insects. Bucket material, shape, and color may vary across experiments but must be held constant within a single sampling effort and matched to the objectives of data collection. #### **Procedure** 1. **Sterilize.** Clean each bucket with a 10% bleach solution and rinse twice with water. To avoid cross-contamination, sterilize on-site in the field or seal buckets during transport from the disinfection site. 2. **Place.** Position the four buckets around the centroid of the plot, approximately 10 m apart. Stabilize so they cannot fall over: brace with sticks on sloping ground; weight with rocks where grazing animals are present. Maintain consistent placement across fields (always in the open, or always near bushes) — comparability between sites matters more than any single placement choice. 3. **Fill.** Add approximately 1 L of filtered water to each bucket. Place a stick taken from the surrounding area into the water so that any insects falling in can escape; we want their eDNA, not the insects themselves. 4. **Expose for 72 hours.** Leave undisturbed. 5. **Observe.** Return after 72 hours and record for each bucket: whether it rained during the exposure period, the fill level, and the presence of any living or dead organisms. Photograph each bucket. 6. **Composite.** Wearing gloves and a face mask, use the sterilized 1 L bottle to measure one litre from each bucket and combine all four into one of the remaining buckets at the site. 7. **Filter and store.** Filter the composite sample using the SimplexDNA kit. The kit operates manually (syringe-driven), is intuitive in the field, and stores the filter in a buffer that preserves DNA at room temperature — a significant practical advantage where refrigeration is unavailable. Filterable volume varies with debris load; the filter clogs progressively and may require considerable force toward the end of filtration. #### **Figure X. Water bucket protocol, colored buckets** --- # Agent Instructions: Querying This Documentation If you need additional information that is not directly available in this page, you can query the documentation dynamically by asking a question. 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