A10. Lamprey ammocoete sampler

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Bottom sampler sketch. Courtesy of INRAE/OFB.

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1. Objectives

Especially designed for young stage lampreys, this device allows to:

  • Sample the 0+ ammocoetes larvae (electrofishing is much less effective on this stage) and know the recruitment of the year
  • Find the growth place of Lampetra fluviatilis and Lampetra planeri, and sometimes sea lamprey (Petromyzon marinus)
  • Avoid electrofishing which requires more resources
  • Know the distribution and population characteristics (abundance, density, structure spatial distribution) that can lead to conservations measures
  • Evaluate impact of a water stream perturbation (cow trampling in the stream)
  • Produce a chronological study on control sites

2. Method summary

Young lampreys are buried in the substrate in the margin of streams and river, and are easily catchable (Lasne and Sabatié, 2012). They prefer sand and fine sediment (silt) with vegetation debris but can be found on detritus covering coarse substrate, in submerged tree roots, and several places of fine sediments accumulation (between vegetation or coarse granulometry), typically with low water velocity.

This technique has been invented by Lasne et al., (2010) as an alternative to electrofishing protocols like quadrat based, multi-run depletion, or calibration (Harvey and Cowx, 2003; JNCC, 2015), which are less effective on the smaller 0+ larvae stage. Others methods also exist, but are less employed (Moser, Butzerin and Dey, 2007; Taverny et al., 2012).

The method is based on a substrate sampling (15 cm deep) collected in a rectangular dredge (30*40*55cm) and filtered through a thin net (1-2mm mesh) to remove clay and silt and collect substrate and ammocoetes. The mesh content is then put in a larger sieve to be sorted, ammocoetes are transferred in a water bucket with anesthetic. The number of ammocoetes and their biometrical characteristics are recorded, whilst substrate and habitat features are noted.

The bottom sampler allows sampling of the three lamprey species. It has shown higher levels of capture of small individuals, compared to electrofishing.

As the number of ammocoetes can be highly variable between samples, a minimum of 30 samples is required to get reliable data on population structure. Species identification can be done morphologically after ammocoetes have reached a given stage or by DNA analysis that allows identification of the separate two genus (Petromyzon/Lampetra) and in a near future separate Lampetra planeri and Lampetra fluviatilis (Lasne, 2020).

3. Advantages

  • This method is standardized and impacts a very restricted area of the river bed
  • Can be done by 2 persons minimum but 3 or 4 is more comfortable and efficient. Low average of 2 sites with 30 samples each day by 2-3 persons. This can be lowered if only aiming for occupancy data
  • Light equipment
  • Useful when there is no fish migration control device (e.g. video-counting system) or on large water basin (mainly concerning Sea lamprey)
  • Does not relying on a specific time period as ammocoete are always in the substrate, although sampling in autumn allows sampling of 0+ and to evaluate reproduction.
  • Provide higher abundance no size bias and is more suitable for smaller lampreys than electrofishing, which is not effective in autumn for <3cm sea lamprey
  • Considers the heterogeneity of ammocoetes distribution in streams. Multiple samples at the low surface provide more accurate results than few big samples
  • Convenient and suitable for small streams

4. Disadvantages

  • Only suitable for shallow waters (40-50 cm depth); beyond that the water enters over the frame
  • Need to sample 0+ between August and October; they are too small before
  • For sea lamprey, adult nesting sites don’t always guarantee the presence of ammocoetes; they can live downstream
  • Lack of data on ammocoete of sea lamprey capture. This may be due to growth taking place deeper in the substrate (Lasne, 2020. com. pers.).
  • No data on efficiency is yet available to compare the “vacuum” method and the “bottom sampler” method
  • Biometry can be the longest step, according to the number of ammocoetes captured

5. Recommendations for method application

  • Know your river basin and the potential sampling area before
  • Be able to identify species and stages, as well as other features such as abundance, population structures etc... which are related to species identification. This can be tricky between the three species
  • Know what specific material is needed, according to protocol.
  • Watch out for depth before choosing the sampling site as water shouldn’t pass over the rectangular dredge
  • Know the life cycle of the species in your area to select the optimal time to catch the largest range of larval class size including 0+ stage, usually between July and October
  • Sample from downstream to upstream to avoid alluvial sediment which may interfere with future samples
  • Concentrate the sampling effort on both sub-optimal and optimal habitats (refer to the field sheet in references)
  • Avoid multiple samples at the same location, once a year maximum

6. Cost

Total equipment cost is below £200. Two or preferably three people are required for the job and can easily do two sites of 30 samples per day.

7. Protocol and data analysis

The absence of ammocoetes in the stream doesn’t mean that adults aren’t visiting the stream, so be careful of misinterpretation. The reasons might be: the larvae survival rate, adult accessibility to the upper stream, or ammocoetes leaving downstream because of hydrology or to find a suitable habitat for growth.

Assessment of population structure and distribution is used to understand population distribution, growth, birth rate and location of nesting areas. Spatial data includes the total abundance on the catchment, stream or station, and mean abundances, as well as the presence of the different stages and their distribution along the catchment or stream habitats.

It is important to record environmental variables like substrate, water depth, velocity, hydromorphology, optimal and sub-optimal habitat surface, and riverbank cover.

Protocols are available at:

Field sheet at the end of this document :

8. Acknowledgement and associated authors

Emilien LASNE - INRAE/OFB - UMR ESE - RENNES

9. Further reading

  • Harvey, J. and Cowx, I. (2003) ‘Monitoring the river, brook and sea lamprey, Lampetra fluviatilis, L. planeriand Petromyzon marinus’, Conserving Natura 2000 Rivers Ecology Series, 5(5), p. 35.
  • JNCC (2015) Common standards Monitoring Guidance for Freshwater fauna.
  • Lasne, E. et al. (2010) ‘A new sampling technique for larval lamprey population assessment in small river catchments’, Fisheries Research. Elsevier B.V., 106(1), pp. 22–26. doi: 10.1016/j.fishres.2010.06.011.
  • Lasne, E. and Sabatié, R. (2012) Méthodologie d’échantillonnage des ammocètes. Extrait du rapport « Flux migratoires et indices d’abondance des populations de lamproies du Scorff, de l’Oir et de la Bresle ».
  • Moser, M. L., Butzerin, J. A. M. and Dey, D. B. (2007) ‘Capture and collection of lampreys: The state of the science’, Reviews in Fish Biology and Fisheries, 17(1), pp. 45–56. doi: 10.1007/s11160-006-9037-3.
  • Taverny, C. et al. (2012) ‘From shallow to deep waters: Habitats used by larval lampreys (genus Petromyzon and Lampetra) over a western European basin’, Ecology of Freshwater Fish, 21(1), pp. 87–99. doi: 10.1111/j.1600-0633.2011.00526.x.