Jul 01, 2026
OceanOmics Centre: 12S, 16S and COI Metabarcoding of Marine Vertebrates
- Laura Missen1,
- Sang Huynh2,1,
- John Blinco3,
- Marcelle Ayad4,
- Georgia Nester1,
- Ebony Thorpe1,
- Anna Depiazzi1,
- Adrianne Doran1,
- OceanOmics UWA1
- 1Minderoo OceanOmics Centre at UWA;
- 2La Trobe University;
- 3Australian Centre for RNA Therapeutics in Cancer;
- 4Minderoo Foundation
External link: https://www.uwa.edu.au/oceans-institute/partnerships/minderoo-oceanomics-centre-at-uwa
Protocol Citation: Laura Missen, Sang Huynh, John Blinco, Marcelle Ayad, Georgia Nester, Ebony Thorpe, Anna Depiazzi, Adrianne Doran, OceanOmics UWA 2026. OceanOmics Centre: 12S, 16S and COI Metabarcoding of Marine Vertebrates . protocols.io https://dx.doi.org/10.17504/protocols.io.rm7vze752vx1/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: January 26, 2026
Last Modified: July 01, 2026
Protocol Integer ID: 241598
Keywords: eDNA, metabarcoding, 12S, 16S, Marine vertebrates, eDNA metabarcoding, automation, biomek i7, Echo 525, environmental DNA, OceanOmics, marine eDNA, COI, biodiversity , marine biodiversity
Abstract
This protocol describes the OceanOmics Centre workflow for generating indexed amplicons for environmental DNA (eDNA) metabarcoding of marine vertebrates, targeting the 16S rRNA, 12S rRNA, and COI gene regions. Samples are amplified with unique combinatorial dual indexes to enable high-throughput multiplexing on Illumina platforms. The workflow has been validated for five metabarcoding assays: 16S-Fish/D, MiFish-U/E2 (multiplexed), MarVer1, and COI-Leray.
The protocol covers the full process from qPCR through to post-qPCR quality control, pooling, and normalisation. Liquid handling is partially automated using the Biomek i7 workstation (Beckman Coulter) and Echo 525 Liquid Handler (LabCyte), with qPCR performed on a Roche LightCycler480 (LC480) using 384-well plates. Associated R scripts for plate mapping and QC, and Biomek scripts for liquid handling, are available in the OceanOmics GitHub repository.
Guidelines
Standard lab set-up:
- Ensure competency and training requirements are met.
- Ensure the laboratory is appropriately configured with separation of pre- and post- PCR rooms. Amplified PCR products should never come in contact with equipment used for non-amplified DNA.
- Wear gloves and a lab coat.
- Use tips with filters to avoid contamination.
- Wipe workspace with 10% bleach, leave to decontaminate for 10 minutes before wiping with ddH20.
- Sterilise all equipment (vortex, minispin, pen markers, outside of tip boxes, pipettes, reservoirs, plates) before use.
- UV pipettes, racks, tubes (lids open), plates and reservoirs for 15 minutes before use.
- Wipe the inside of the centrifuge and rotor with 70% ethanol.
- Decontaminate Biomek deck before use with DNA erase and 70% ethanol. Tighten mandrels, empty waste and check liquid input and waste levels.
- Ensure all reagents are aliquoted in appropriate amounts and stored according to manufacturers' recommendations. Never pipette directly from reagent stocks. Bring all reagents to room temperature before use.
- All waste must be disposed of following institutional biosafety regulations.
- Include a Negative Template Control (NTC) and Positive Control (ITC) on every plate.
- Mastermix should be prepared in low light conditions.
Materials
Equipment
Equipment
Biomek i7 Automated Workstation
NAME
Automated Liquid Handlers
TYPE
Beckman Coulter
BRAND
B87585
SKU
LINK
Biomek i7 Hybrid (MC + Span-8) with Enclosure
SPECIFICATIONS
Equipment
Echo 525 Liquid Handler
NAME
Labcyte
BRAND
001-10080
SKU
LINK
- PCR laminar flow hood
- Centrifuge with rotor for 1.5 mL tubes and 96-well plates
- Roche LightCycler 480
- Biomek i7 24-position tube rack holder
- Biomek i7 tube rack inserts (11 mm, white)
- UV sterilisation cabinet
- Beckman Coulter rack for reservoirs
- Heat Sealer
Consumables
- Eppendorf twin.tec PCR Plate, 96, skirted, 250 µL
- Eppendorf cap strips, 8-strip, domed
- 50 mL conical tubes
- Eppendorf 1.5 mL DNA LoBind tubes
- Bio-Rad Hard-Shell 384-well PCR plates, clear/white, barcoded
- Axygen CyclerSeal PCR sealing film
- Axygen Ultra Clear Pressure Sensitive sealing film
- Eppendorf heat sealing film
- Beckman Coulter Biomek i-Series 50 µL filtered tips
- Beckman Coulter Biomek i-Series 1025 µL filtered tips
- Beckman Coulter Reservoir, 40 mL
- Fisherbrand 96-well deep well polypropylene plates
- Beckman Coulter Echo qualified 384-well polypropylene source microplate
- Kimwipes
Reagents
- 10% bleach
- 70% ethanol
- ddH2O
- LookOut DNA Erase
AmpliTaq Gold™ 360 Master MixThermo FisherCatalog #4398901
Bovine Serum Albumin (20 mg/ml)Thermo FisherCatalog #B14
Nuclease-free Water (not DEPC-Treated)Thermo FisherCatalog #AM9937
SYBR Green (10000x)Thermo Fisher ScientificCatalog #S7563
Safety warnings
This protocol involves the use of potentially hazardous biological materials, chemicals (including ethanol and enzymatic reagents), and electrical laboratory equipment. Always wear appropriate personal protective equipment (PPE), including a lab coat, gloves, and eye protection. Be familiar with the relevant Safety Data Sheets (SDS) and laboratory risk assessments before commencing work. Follow institutional safety guidelines and dispose of waste in accordance with local regulations. Exercise caution when handling sharp instruments, thermal cyclers, and high-speed centrifuges to prevent injury or equipment damage.
Relevant SDS documents: 
AmpliTaq Gold 360 SDS.pdf105.8KB BSA SDS.pdf162.6KB SYBR SDS.pdf110.2KB
AmpliTaq Gold 360 SDS.pdf105.8KB BSA SDS.pdf162.6KB SYBR SDS.pdf110.2KB All scripts were developed in house on the Beckman Coulter Biomek i7 automated workstation, and the LabCyte Echo 525. They have been optimised for specific consumables and reagents, any substitutions or deviations may affect performance.
All R-scripts were developed specifically to handle data from the Roche Diagnostics LC480. If using an alternative qPCR instrument, the scripts will need to be modified.
Before start
Prepare working environment as per the guidelines.
Download all relevant R-scripts for qPCR set-up and pooling from GitHub: https://github.com/Minderoo-OceanOmics-Centre-UWA/qPCR_setup_pooling
Download all relevant Biomek i7 and Echo 525 Protocols: https://github.com/Minderoo-OceanOmics-Centre-UWA/OceanOmics_Biomeki7_Automated_Methods
Create Plate Templates
This step assumes DNA extracts are supplied in tubes. It walks through using an R script to generate a plate map to aliquot DNA from tubes into a 96-well DNA input plate, a plate map for the 384-well qPCR plate, and the assignment of unique combinatorial index sequences to each sample during qPCR.
Note
During qPCR, each sample is run in triplicate, occupying positions 1–3 within its quadrant, as shown below. Position 4 of each quadrant is left empty during qPCR and reserved for pooling the triplicate reactions afterwards.
In the attached Input_Template, enter all required samples in the Metadata tab and assign each one to its corresponding project. Do not edit any other tab, these contain the primer and index sequences for all assays.
Input_Template.xlsx38.8KB NOTES:
- Index combinations can only be used once per assay within a single sequencing run. Reusing an index combination on the same run will cause sample reads to be mixed together and cannot be demultiplexed. Sequencing runs must therefore be planned before qPCR to ensure no index combinations overlap.
- Target read depth must also be defined in advance, based on the number of samples going onto the run, to ensure the sequencing run capacity is filled appropriately.
Once all relevant R-scripts for qPCR set-up and pooling have been downloaded from GitHub, follow the README instructions to ensure files are in the correct folder and format.
Run the 'run_meta_to_plates' R-Script. This script takes the completed Input_Template.xlsx as input and generates an output dataframe (df), exported as an Excel file, which maps samples to plate positions and index combinations. The most important tabs are:
- "plates" — DNA template plate map (96-well format), showing the position of each sample on the source DNA plate.
- "big_plates" — qPCR plate map (384-well format), showing the layout of each sample as a triplicate, alongside its assigned indexed primer pair.
- "metadata" — assigns each sample to its index plate position and index sequence.
- "position_df" — assigns sample triplicates to their qPCR plate positions.
NOTE: as a default, this template will reserve the bottom right two quadrants of the plate for a negative control (NTC) and a positive control (ITC).
Plating DNA into a 96-well plate using Biomek i7
Download the template below and fill in the sample ID (column F), listing samples in the same order as the metadata tab generated in Step 4. Do not edit any other column. Save as a csv file.
Important: Before proceeding, check that the DNA well positions in this template match the well positions generated by the R-script above. Any mismatch will cause samples to be assigned the wrong index combination.
TEMPLATEbiomek_eDNA_Plating_96tubes.csv2.3KB Upload this .csv file to the Biomek PC.
Download and open the following protocol on the Biomek PCR: eDNA_plating_byCSV_JB_SH_v04
Ensure the correct instrument files have also been uploaded to the Biomek PC.
Insert white tube inserts into the Biomek tube rack holder in all 24-positions.
Insert DNA extract tubes into the tube rack according to the positions listed on the .csv file. Ensure the lid is open and held back in position and the tube is fully inserted. Avoid touching the inside of the lid.
NOTE: it is important to decontaminate the rack and inserts after each use with 10% bleach.
Home the Biomek, then run the protocol and follow the prompts. Do not change any values.
Select the desired .csv file in the file opener window.
Method Launcher will open once guided set-up has been executed. Follow the guidance to position the labware on the deck. An example of a deck layout is shown below:
In this protocol, 25 µL of eDNA extract is dispensed into the plate. Freeze the remaining eDNA extract tubes at -20°C.
Safe stopping point.
Once complete, seal the eDNA plate with 8-strip domed caps and store at 4°C until required for processing.
Empty tip bins on the biomek, and decontaminate all surfaces with DNAErase, wait until fully dry before wiping with 70% ethanol.
Mastermix Preparation
This section covers the mastermix recipe and dispensing method onto 384-well plates using the Biomek i7.
Once the mastermix has been prepared, it is first dispensed by the Biomek into a 96-well deep well plate. Manual intervention is then required: remove the plate from the deck, centrifuge it, and return it to the deck. The Biomek then dispenses the Mastermix from the deep-well plate into the 384-well plate.
Wipe reagent hood with 10% bleach and leave to decontaminate for 10 minutes before wiping with distilled water, then UV for 20 minutes.
NOTE: reagent handling must be kept separate from sample handling.
Mastermix Recipe:
Using the attached calculator (qPCR_Calculator.xlsx), determine the volume of each reagent required for the total number of samples. Enter the number of qPCR plates and the number of assays into the blue cells. The orange cells will then display the volumes required for the mastermix.
qPCR_Calculator.xlsx25.9KB NOTE: the total volume will highlight green if it falls within the 40 mL limit — this limit reflects the maximum capacity of the trough used to hold the mastermix on the Biomek. If it highlights red, reduce the number of plates or assays. A maximum of 9x 384-well plates can be prepared in one batch.
Combine the AmpliTaq, BSA, SYBR Green, and water together in a size-appropriate tube to create the mastermix. Invert gently 5x to mix. Do not vortex as this will create bubbles.
NOTE: SYBR Green is light-sensitive. Keep the mastermix covered/protected from light and minimise exposure throughout preparation and handling.
Open the Biomek 5 software, Home the instrument then open the following protocol: qPCR_MM_setup_JBSHv06
Once guided set-up has been executed, Method Launcher will open. Follow the on-screen guidance to position the labware on the deck, as shown in the example layout below:
NOTES:
- All plates should be positioned with A1 in the top left corner.
- Labware includes Fisherbrand 96-well deep well polypropylene plates (interim plate) and Bio-Rad Hard-Shell 384-well PCR plates, clear/white, barcoded (final plate).
Pour the mastermix into a 40 mL Beckman Coulter reservoir then place into the MMRes ALP on the Biomek deck.
Start the protocol and follow the prompts to fill the number of plates (maximum 9). All other values should be kept at 0.
When prompted, designate which Tips8Span (Beckman Coulter Biomek i-Series 1025 µL filtered tips) are available by highlighting where available in blue.
The Biomek will pause for pre-run checks, select 'OK' once you have checked all labware is in the correct position and orientation. Do not rely on the instrument cameras to check deck layout, ignore errors and check manually.
There is a pause step after the Biomek has dispensed into the 96-well deep well plate. During this pause, remove the plate, seal, and spin-down at 1000 x g for 1 minute to remove air bubbles. Remove seal and place back into the same position on the Biomek deck, then select 'Start'.
Once complete, remove all plates from the deck. The 384-well plate now contains the Mastermix and can be sealed with an Axygen Ultra Clear Pressure Sensitive seal; cover it with aluminium foil to protect from light. The 96-well deep-well plate can be discarded.
Safe stopping point.
If proceeding with qPCR the same day, store at 4°C until required.
Alternatively, plates can be stored at -20°C for up to one month. Ensure they are protected from light.
Transfer tagged primers to qPCR plate
This section describes how tagged primers are transferred from Echo Source Plates (Beckman Coulter Echo-qualified 384-well polypropylene source microplates) into the pre-prepared 384-well plate containing mastermix.
You will need:
- Pre-prepared 384-well plate containing mastermix.
- Tagged primers prepared in Echo Source Plates (Beckman Coulter Echo qualified 384-well polypropylene source microplate). Prepare 60 µL of each 60 µM indexed primer in the configurations below:
16S_Echo_Source_Plates.xlsx46.5KB MiFishU_Echo_Source_Plates.xlsx40.4KB MiFishE2_Echo_Source_Plates.xlsx39.3KB MarVer1_Echo_Source_Plates.xlsx39.3KB COI_Echo_Source_Plates.xlsx41.1KB NOTE: ensure all indexed primers are diluted to 60 µM before use.
Download the following protocols for Echo 525: index_plating.epr (for single assays) and index_plating_multiplexing.epr (for dual-plexed assays).
NOTE: MiFish-U and MiFish-E2 can be multiplexed together
Thaw the required plates at room temperature for 30 minutes then centrifuge at 2000 x g for 1 minute.
Using the Echo Software, survey the plates, ensuring that the volume in each well is between 12 µL and 65 µL. If the value is too low, prepare a new plate, if the value is too high, there may be air bubbles or droplets on the plate. Spin-down and survey again.
NOTES:
- Wipe the bottom of the plates with a Kimwipe before loading onto the Echo.
- See Echo 525 manual for detailed instructions on how to survey plates and run protocols.
For standard assays, run the 'index_plating' protocol on the Echo 525. For multiplexing assays, run the 'index_plating_multiplexing' protocol on the Echo 525.
Check the 'output_dataframe' R file to see which index plate the qPCR plate is associated with.
NOTE: for multiplexed assays (MiFish-U/E2), each qPCR plate will undergo two transfers (one for each assay).
Once indexes have been transferred, re-seal the index plates using a heat sealer and place back into the freezer. For qPCR plates, proceed immediately to the next step.
NOTE: the index plates should be freeze/thawed a maximum of 10 times, keep a record to track this.
Transfer DNA template and controls to qPCR plate
This section describes how 2 µL DNA (from the 96-well plate) is transferred in triplicate into the pre-prepared 384-well plate containing mastermix and tagged primers.
Bring the previously prepared 96-well DNA plate to room temperature then spin-down at 1000 x g for 1 minute.
Open the Biomek 5 software, Home the instrument and then open the following protocol: qPCR_Sample_transfer_JBv06
Start the protocol and follow the prompts to the fill the number of plates, no other variables should be changed.
Method Launcher will open once guided set-up has been executed. Follow the guidance to position the labware on the deck. An example of a deck layout is shown below:
NOTE: all plates should be positioned with A1 in the top left corner.
The Biomek will pause for pre-run checks, press 'OK' once you have checked all labware is in the correct position. Do not rely on the instrument cameras to check deck layout, ignore errors and check manually.
For this method, you will need:
- DNA input plate (96-well plate)
- 384-well plate containing mastermix and tagged primers
- Beckman Coulter Biomek i-Series 50 µL filtered tips
Once complete, place qPCR plates into a PCR-hood for the addition of controls.
Seal the DNA plates with 8-strip domed caps. If using the samples again, place into the fridge. If the samples are not required again, place at -20°C.
Dispense 2 µL of nuclease-free water into the corresponding NTC wells of the qPCR plate. Pipette to mix.
Dispense 2 µL of a positive control (e.g. 0.1 ng/µL Zebrafish genomic DNA) into the corresponding ITC wells of the qPCR plate. Pipette to mix.
NOTE: use a positive control that cannot be present in your samples, so for marine environments a freshwater fish is used.
Seal the plate with an Axygen CyclerSeal then spin-down at 2000 x g for 1 minute.
qPCR Conditions
qPCR cycling conditions vary by assay. Run each qPCR plate on the LightCycler480 (LC480) according to the conditions listed below for its corresponding assay:
QC of qPCR results
This section describes the QC method to perform once qPCR has finished on the LightCycler480 (LC480) system. It covers how to assess whether samples and/or replicates pass the QC thresholds (discussed further below), and how to calculate normalisation volumes to account for differences in amplicon amplification between samples.
Download all EPF, Cp, and Tm files from the LC480 for all qPCR plates. The following naming conventions are required for the files to be read by the R-script:
yymmdd_projectid_assay_platenumber_metric
e.g. 260630_OcOm001_MiFishU_Plate1_EPF
Run the 'run_concat_qPCR_data' R script. This will concatenate all the data files into one data frame for each plate. These files are then used as the input for the next script.
NOTE: follow the README instructions on GitHub to ensure files are in the correct folder and format.
Run the 'run_plates_to_pooling' R script. This script collates all LC480 output files, identifies failed replicates and failed samples, and calculates relative pooling volumes based on each sample's EPF value. It then generates a .csv file, which is used as the input for the Biomek i7 amplicon pooling protocol (described in the section below).
Partway through, the script generates a file called 'rnxs_to_check.csv'. This file flags samples that require a manual QC check before the script can continue. For example, a sample might have a Cp value below 40, but its Tm doesn't match the expected amplicon size — in this case, you would manually change that sample's status from KEEP to DISCARD.
Once you've reviewed the file, save it as a .csv named 'rxns_checked.csv' in the output directory, then continue running the R script.
Use the table below as a guide for these manual QC checks.
NOTES:
- A failed sample is defined as a sample in which 2 or more replicates fail to meet QC criteria. These samples will not be taken through to pooling.
- If one replicate fails, this replicate is manually discarded from the plate and then the other two replicates are taken through to sequencing.
- All controls are taken through to sequencing.
This script relatively pools samples using EPF values, so the higher the EPF, the lower the volume pooled. This is the first step of normalisation to ensure samples receive an equal number of reads during sequencing.
Check the 'biomek_pooling_workbook.csv' to ensure that each plate has two tube positions assigned (e.g. A1 for samples and B1 for controls), and that there are also tube positions assigned for NTCs (e.g. C1) and ITCs (e.g D1).
An example is attached:
biomek_pooling_workbook_example.csv10.9KB Amplicon Pooling
This section describes the amplicon pooling process, where indexed PCR products are combined into a 1.5mL DNA LoBind tube pool. Once QC is complete, amplicons are relatively pooled based on their EPF value, using the Biomek i7 workstation. Each plate produces 2x pools: one for samples and one for controls. Each assay additionally produces 2x control pools (NTC and ITC). Samples with a higher EPF value are pooled at a lower volume, and vice versa, ensuring each sample is represented at a more even concentration in the final pool. This relative pooling is the first stage of normalisation prior to library quantification and final pooling for sequencing.
Remove all qPCR plates from the freezer and allow to thaw at room temperature.
Centrifuge plates at 2000 x g for 5 minutes to ensure all droplets are removed from the plate seal.
Use the 'reps_to_discard.csv.' output file to identify which replicates to DISCARD from the plates. Aspirate and discard 15 µL of all relevant replicates.
Download the 'biomek_pooling_workbook.csv' output file onto a USB and import to the Biomek i7 PC.
Open the Biomek 5 software, Home the instrument then open the following protocol: qPCR_Pooling_JBv12_LowBind
Start the protocol. The protocol allows you to choose whether to run triplicate pooling, cherry picking, or both. To run the triplicate pooling enter the value 'True' for 'Do384pool' when prompted. To run cherry picking, enter the value 'True' for 'DoCherryPick' when prompted. Enter 'False' if not required.
Note
Do384pool: pools triplicate reactions into the reserved fourth well of each quadrant (equal volumes)
DoCherryPick: relatively pools samples by EPF (unequal volumes)
When prompted, select the desired file ('biomek_pooling_workbook.csv.').
When prompted, position labelled 1.5 mL DNA LoBind tubes in the tube rack in the positions shown on the prompt. For example:
NOTES:
- Ensure the lids are fully snapped back into position, and the tube is inserted fully. Any deviations could cause the instrument to crash.
- Ensure tubes are labelled with the plate number, assay, sample/control, and that they are in the correct position in the tube rack. Use the 'biomek_pooling_workbook.csv' to guide this.
Method Launcher will open once guided set-up has been executed. Follow the guidance to position the labware on the deck. An example of a deck layout is shown below:
A maximum of 4 qPCR plates can be pooled at a time. These should be positioned as follows:
(Optional) A partial tip box can be used if there is one already open. If using a partial tip box, designate the location of the tips by highlighting where they are available. There must be a minimum of 8 tips in the box.
The Biomek will pause for pre-run checks, press 'OK' once you have checked all labware is in the correct position. Do not rely on the instrument cameras to check deck layout, ignore errors and check manually.
The instrument will automatically pause after running the triplicate pooling step. Seal and spin-down qPCR plates then place back into the correct positions on the Biomek deck and press 'OK'. This will start the cherry picking step.
Once complete, amplicons are relatively pooled based on their EPF. There should be 2x pools per plate (Samples and Controls), for each assay there will be an extra 2x control pools (NTC and ITC).
Safe stopping point.
Amplicon pools can be stored at 4°C short-term, or -20°C long-term. Avoid freeze/thaw cycles.
Pools must then be cleaned and normalised before library preparation, proceed to one of the following protocols:
OR
OceanOmics Centre: eDNA Amplicon Clean-Up and Pooling (Automated)
Protocol references
MiFish-U:
Miya, M., Sato, Y., Fukunaga, T., Sado, T., Poulsen, J., Sato, K., Minamoto, T., Yamamoto, S., Yamanaka, H., & Araki, H. (2015). MiFish, a set of universal PCR primers for metabarcoding environmental DNA from fishes: detection of more than 230 subtropical marine species. Royal Society open science, 2(7), 150088.
MiFish-E2:
Miya, M. a. S., T. (2019). Multiple species detection using MiFish primers. Pages 55–92 in
Environmental DNA sampling and experimental manual version 2.1. Ed. by eDNA Methods
Standardization Committee, The eDNA Society, Otsu, Japan.
MarVer1:
Valsecchi, E., Bylemans, J., Goodman, S. J., Lombardi, R., Carr, I., Castellano, L., Galimberti, A., & Galli, P. (2020). Novel universal primers for metabarcoding environmental DNA surveys of marine mammals and other marine vertebrates. Environmental DNA, 2(4), 460-476.
16S-Fish/D:
Berry, T. E., Osterrieder, S. K., Murray, D. C., Coghlan, M. L., Richardson, A. J., Grealy, A. K., Stat, M., Bejder, L., & Bunce, M. (2017). DNA metabarcoding for diet analysis and biodiversity: A case study using the endangered Australian sea lion (Neophoca cinerea). Ecology and evolution, 7(14), 5435-5453.
Deagle, B. E., Gales, N. J., Evans, K., Jarman, S. N., Robinson, S., Trebilco, R., & Hindell, M. A. (2007). Studying seabird diet through genetic analysis of faeces: a case study on macaroni penguins (Eudyptes chrysolophus). PloS one, 2(9), e831.
COI-Leray:
Leray, M., J. Y. Yang, C. P. Meyer, et al. 2013. A New Versatile Primer Set Targeting a Short Fragment of the Mitochondrial COI Region for Metabarcoding Metazoan Diversity: Application for Characterizing Coral Reef Fish Gut Contents. Frontiers in Zoology 10: 34.