A study of phytoliths produced by selected native plant taxa commonly used by Great Basin Native Americans

Abstract

Reference collections of phytoliths from plant taxa commonly used by the ancient inhabitants of an archaeological site are critical to researchers conducting analyses on these microbotanical remains. Phytoliths recovered from the site are compared to those in reference collections to make inferences about which taxa were being used, as well as where, why and how. This study presents one of the first reference collections of phytoliths from selected taxa likely to have been used by Native American peoples living in the Great Basin area. The results are presented as a list of taxa which we sampled, and that are known to have been used by these tribes, such as the Shoshone and Ute, with illustrations of the range of phytolith morphotypes produced by each taxon. Our hope is that this tool will prove to be a ready and valuable resource for any researchers conducting phytolith analysis at Great Basin Native American sites.

Introduction

Opal phytoliths are solid inorganic structures which form in many plant taxa as a plant takes up monosilicic acid, Si(OH₄), through its roots and deposits it as silica in and around the plant cells (Pearsall et al. 1995, p. 184). In the plant cell walls and lumens of some taxa, the silica forms a solid deposit that takes the shape of the cell or space in which it formed. In some cases, this shaped silica, or phytolith, can be taxonomically significant (Pearsall et al. 1995, p. 184; Ball et al. 2016).

When a plant’s organic components are destroyed through processes such as decay, burning, digestion or grinding, any phytoliths contained there are released into the surrounding environment, thus becoming microfossils of the plant (Ball et al. 1999, p. 1615; Piperno 2006). These microfossils can then be recovered from several different archaeological contexts such as soil, coprolites, dental calculus, stomach contents, residue on artifacts, or lake cores (Berlin et al. 2003, p. 115; Piperno 2006, pp. 81–86). Such phytolith studies have added to both prehistoric and present day environmental and ethnobotanical reconstruction (Ball et al. 2015).

Compared to other plant microremains, opal phytoliths have three characteristics that can make them especially useful for archaeobotanical investigations. First, due to their inorganic nature, phytoliths preserve better in some environments such as highly oxidized soils that typically destroy the organic components of other microbotanical remains (Pearsall 1989, p. 254). Second, unlike plant disseminules such as spores, pollen and seeds that are primarily produced during specific seasons or stages of development, some types of phytoliths, such as those produced in the leaves of many taxa, can be produced in a plant throughout its entire life cycle. Finally, some types of phytoliths can be produced in plant tissues and organs that do not produce other forms of microbotanical remains (Ball et al. 2015, p. 11).

Any analysis of archaeological phytoliths recovered from an excavation relies upon the researcher’s ability to distinguish between the different taxa that may have produced them. As a first step towards that end, researchers typically assemble reference collections of phytoliths produced by the plant taxa that may have been used by the ancient inhabitants of the site, as well as those of the native and non-native vegetation in the area. Archaeological phytoliths recovered from the site are then compared to those in the reference collections to make inferences about which taxa were being used, as well as where, how and why. This study presents an initial or baseline survey of phytolith types produced by selected taxa likely to have been used by Native American peoples, such as the Shoshone, Ute and Southern Paiute who were predominantly in Nevada and Utah, and different tribes in Oregon and western Nevada (ESM 1). The list of taxa which we sampled has illustrations of the phytolith morphotypes that we observed in a single sample from each taxon, with botanical nomenclature following Welsh et al. (2008) and The Plant List (2019). Our list of morphotypes described should be viewed as illustrative rather than exhaustive or diagnostic for any given taxon, as this is an initial survey. Still, it provides an important first step and baseline that we hope will invite further research and the development of robust reference collections based on replicate analyses of many samples.

Methods

We made a list of 160 plant species native to the Great Basin that have documented ethnographic uses by the Shoshone tribe including the Goshute, the Southern Paiute and the Ute tribes (Pearce 2017). We collected various tissue samples from a single example of 52 of these listed taxa from botanical gardens, nurseries, herbariums and wildlife and recreation areas in both Utah and Salt Lake Counties and in the rest of the state of Utah (ESM 2). All non-herbarium samples were collected during the spring and summer of 2015; all herbarium samples were collected during the winter of 2014. We failed to record voucher numbers for the specimens, but intend to do so in future studies and recommend this to other researchers. Tissue sample sizes ranged from a few leaves to an entire plant depending on access to the plant and sampling permission limits.

To prepare the material we used the acid digestion methods described by Portillo et al. (2006) to extract any phytoliths from our samples of the 52 taxa. Various plant tissues or organs were processed separately to extract the phytoliths from each. For example, because the berries and leaves of Shepherdia canadensis (L.) Nutt. have different documented ethnographic uses, we processed both of them separately for phytoliths.

We found that some plant material was more difficult to digest than usual using the acid digestion method, so occasionally extra grinding and drying of the plant material was required before digestion, followed by repeated acid treatments to remove all the organic content. For example, the inflorescences of Achillea millefolium L. required two acid treatments for complete digestion, while the leaves of S. canadensis required three. We assumed that multiple acid treatments would not affect the observed phytolith production index (PI), but further tests should be conducted to confirm this assumption.

We also found it helpful to sonicate the plant material in a mild cleaning solution such as Teepol before acid treatment to remove contaminants such as terrestrial diatoms, dust and other debris that may have adhered to the outer surfaces of the sample, for 5 min using a Mettler Cavitator ultrasonic cleaner. Then the plant material was rinsed by placing it in a clean beaker and sonicating for an additional 5 min in distilled water. Any required grinding or drying followed the sonication before beginning the acid treatment. Sonication did not, we assume, affect the relative abundances of phytoliths observed, but this assumption should be tested in the future.

The phytoliths extracted from the samples were mounted on glass slides under a cover slip using Permount, for light microscope analysis. We used either a Zeiss Axiovert 135 or a Nikon Optiphot 2 light microscope with an attached Infinity 2 camera at magnifications of ×100, ×200, or ×400 to identify and collect images of the phytolith morphotypes produced by each taxon. All morphotypes are described using the International Code for Phytolith Nomenclature 2.0 (ICPN 2.0) (Neumann et al. 2019) and the International Code for Phytolith Nomenclature 1.0 (ICPN 1.0) (Madella et al. 2005). We primarily relied upon ICPN 2.0, which will be published soon and replaces ICPN 1.0.

Results

Of the 52 plant species that we analysed (ESM 1), we found that 24 contained identifiable phytolith morphotypes, 21 had less distinctive vascular tissue phytolith types and seven had no phytoliths (Figs. 1, 2, 3, 4, 5, 6, 7, 8, and 9). Again, we note that because we sampled only a single specimen of each taxon, our findings should be considered illustrative rather than exhaustive or diagnostic for the types and numbers of phytoliths produced within the taxa. Further detailed studies of each taxon that include quantified samples of many specimens will likely provide a better range of variability, but we assume the most common morphotypes likely to be produced by our selected taxa are included in this study.

Fig. 1

acute phytoliths—acute crassus granulate: aPoa fendleriana (Steud.) Vasey; acute crassus psilate: bAchillea millefolium L.; cDeschampsia cespitosa (L.) P.Beauv.; dLeymus cinereus (Scribn. & Merr.) Á.Löve; eElymus glaucus; fStipa hymenoides Roem. & Schult.; acute gracile psilate: gAmelanchier alnifolia (Nutt.) Nutt. Ex M.Roem.; hFestuca ovina L.; acute gracile psilate echinate: iLeymus cinereus (Scribn. & Merr.) Á.Löve; acute gracile psilate/granulate: jHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; acute gracile psilate/granulate segmented: kBalsamorhiza sagitta (Pursh) Nutt.; lSolidago canadensis L.; acute gracile striate/granulate: mArtemisia dracunculus L.; acute bulbosis echinate: nLeymus cinereus (Scribn. & Merr.) Á.Löve; oElymus glaucus Buckley; acute bulbosis/psilate segmented: pHeliomeris multiflora Nutt. Scale bars 20 μm, image g scale bar 50 μm

Fig. 2

Articulated epidermal phytoliths—elongate psilate columnar/clavate: aLeymus cinereus (Scribn. & Merr.) Á.Löve; bElymus glaucus Buckley; elongate psilate entire/sinuate: cLeymus cinereus (Scribn. & Merr.) Á.Löve; dElymus glaucus Buckley; eHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; f and gStipa hymenoides Roem. & Schult.; elongate/irregular psilate/striate entire/sinuate: hHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; iRhus aromatica Aiton.; elongate/oblong/irregular psilate entire/sinuate: jAchillea millefolium L.; kArtemisia ludoviciana Nutt; lArtemisia tridentata Nutt; mDeschampsia cespitosa (L.) P.Beauv; nJuniperus communis L; oRosa woodsii Lindl.; p and qSporobolus airoides (Torr.) Torr.; rStipa hymenoides Roem. & Schult.; elongate/polygonal psilate entire: sRosa woodsii Lindl; tShepherdia argentea (Pursh). Nutt. Irregular psilate/granulate sinuate: uSphaeralcea munroana (Douglas ex Lindl.) Spach ex A.Gray; Irregular psilate sinuate: vArtemisia dracunculus L.; wEriogonum umbellatum Torr.; xHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller. Scale bars 20 μm

Fig. 3

Articulated epidermal, irregular and circular/ovate phytoliths—Irregular psilate sinuate: aPrunus virginiana L.; bSporobolus airoides (Torr.) Torr.; Irregular striate sinuate: cArtemisia biennis Willd.; dArtemisia dracunculus L.; eHedysarum boreale Nutt.; Irregular psilate sinuate/velloate/entire: fAmelanchier utahensis Koehne; gHeliomeris multiflora Nutt.; Irregular/circular/ovate striate sinuate/entire: hGutierezia sarothrae (Pursh) Britton & Rusby.; Polygonal psilate entire: iArtemisia dracunculus L. Favose; jEriogonum umbellatum Torr.; Irregular circular/ovate: kCercocarpus ledifolius Nutt. ex Torr. & A.Gray; lEriogonum ovalifolium Nutt.; mHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; nRhus aromatica Aiton; oSolanum jamesii Torr.; Circular/ovate: pArtemisia ludoviciana Nutt; qArtemisia tridentata Nutt; rBalsamhoriza sagittata (Pursh) Nutt; sDeschampsia cespitosa (L.) P.Beauv; tFestuca ovina L.; uGutierezia sarothrae (Pursh) Britton & Rusby; vHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; wPinus edulis Engelm; xPrunus virginiana L.; yRhus aromatica Aiton. Scale bars 20 μm

Fig. 4

Circular/ovate, Papillate, stomata, tracheary pitted and other phytoliths—Circular/ovate: aRosa woodsii Lindl; bSphaeralcea munroana (Douglas ex Lindl.) Spach ex A.Gray. Astrosclerid: cArtemisia biennis Willd.; dGutierezia sarothrae (Pursh) Britton & Rusby.; Lunate granulate: ePinus edulis Engelm.; papillate: fAchillea millefolium L.; gLeymus cinereus (Scribn. & Merr.) Á.Löve; hElymus glaucus Buckley; iPoa fendleriana (Steud.) Vasey. Stomata: jArtemisia dracunculus L.; kSphaeralcea munroana (Douglas ex Lindl.) Spach ex A.Gray.; Umbraculiform striate: lShepherdia argentea (Pursh). Nutt.; tracheary pitted/annulate: mFestuca ovina L.; tracheary pitted curled: nDeschampsia cespitosa (L.) P.Beauv; oFestuca ovina L.; tracheary pitted: pDeschampsia cespitosa (L.) P.Beauv; qFestuca ovina L.; rPoa fendleriana (Steud.) Vasey; sSporobolus airoides (Torr.) Torr.; tStipa hymenoides Roem. & Schult.; tracheary pitted pilate: uAchillea millefolium L. Scale bars 20 μm, image d scale bar 50 μm

Fig. 5

tracheary annulate/helical phytoliths—tracheary annulate/helical: aAchillea millefolium L.; bAmelanchier utahensis Koehne; cArtemisia dracunculus L.; dArtemisia ludoviciana Nutt.; eArtemisia tridentata Nutt.; fBalsamhoriza sagittata (Pursh) Nutt.; gEricameria nauseosa (Pall. ex Pursh) G.I.Nesom & G.I.Baird; hDeschampsia cespitosa (L.) P.Beauv; iLeymus cinereus (Scribn. & Merr.) Á.Löve; jEphedra nevadensis S.Watson; kEphedra viridis Coville; lEriogonum umbellatum Torr.; mGutierezia sarothrae (Pursh) Britton & Rusby; nHolodiscus dumosus (Nutt. ex Torr. & A.Gray) A.Heller; oOpuntia polycantha Haw.; pPrunus virginiana L.; qRhus aromatica Aiton; rRosa woodsii Lindl; sSarcobatus vermiculatus (Hook.) Torr.; tShepherdia argentea (Pursh). Nutt.; uSolanum jamesii Torr.; vSolidago canadensis L. Scale bars 20 μm

Fig. 6

elongate phytoliths—elongate dendritic/dentate: a, bLeymus cinereus (Scribn. & Merr.) Á.Löve; c, dElymus glaucus Buckley; eFestuca ovina L.; fPinus monophyla Torr. & Frém.; elongate entire granulate: gFestuca ovina L.; hGutierezia sarothrae (Pursh) Britton & Rusby; iSporobolus airoides (Torr.) Torr.; jStipa hymenoides Roem. & Schult.; elongate psilate columnar/clavate/sinuate: kLeymus cinereus (Scribn. & Merr.) Á.Löve; lElymus glaucus Buckley; elongate psilate entire: mDeschampsia cespitosa (L.) P.Beauv; nFestuca ovina L.; oPoa fendleriana (Steud.) Vasey; pSporobolus airoides (Torr.) Torr.; qStipa hymenoides Roem. & Schult.; elongate psilate/granulate echinate/baculate: rDeschampsia cespitosa (L.) P.Beauv; sLeymus cinereus (Scribn. & Merr.) Á.Löve; tFestuca ovina L., u, vPoa fendleriana (Steud.) Vasey; w, xSporobolus airoides (Torr.) Torr.; y, zStipa hymenoides Roem. & Schult. Scale bars 20 μm

Fig. 7

spheroid and blocky phytoliths—spheroid ornate/ellipsoidal granulate/plicate: aAchillea millefolium L.; bArtemisia biennis Willd.: cArtemisia dracunculus L.; dBalsamhoriza sagittata (Pursh) Nutt.; eEricameria nauseosa (Pall. ex Pursh) G.I.Nesom & G.I.Baird; fEriogonum ovalifolium Nutt; gGutierezia sarothrae (Pursh) Britton & Rusby; hJuniperus osteosperma (Torr.) Little; iPrunus virginiana L.; jPurshia tridentata (Pursh) DC.; kRhus aromatica Aiton; lRibes aureum Pursh.; mShepherdia canadensis (L.) Nutt; nSolanum jamesii Torr; oSolidago canadensis L.; spheroid ornate/ellipsoidal baculate/pilate: pLeymus cinereus (Scribn. & Merr.) Á.Löve; qElymus glaucus Buckley; blocky/tabular/irregular psilate/granulate: rArtemisia ludoviciana Nutt.; sArtemisia tridentata Nutt.; tAtriplex truncata (Torr.) A.Gray; u, vEricameria nauseosa (Pall. ex Pursh) G.I.Nesom & G.I.Baird; wCrataegus douglasii Lindl.; xEphedra nevadensis S.Watson; yEriogonum umbellatum Torr; zOpuntia polycantha Haw.; aaPurshia mexicana (D.Don) Henr.; bbPurshia tridentata (Pursh) DC. Scale bars 20 μm

Fig. 8

Grass Silica Short Cell Phytoliths (GSSCP)—rondel: a–eDeschampsia cespitosa (L.) P.Beauv; f–hLeymus cinereus (Scribn. & Merr.) Á.Löve; i–lElymus glaucus Buckley; m–pFestuca ovina L.; q–sPoa fendleriana (Steud.) Vasey; tSporobolus airoides (Torr.) Torr.; u, vStipa hymenoides Roem. & Schult.; saddle: wSporobolus airoides (Torr.) Torr.; bilobate/cross/polylobate/elongate: xDeschampsia cespitosa (L.) P.Beauv; yPoa fendleriana (Steud.) Vasey; zSporobolus airoides (Torr.) Torr. Scale bars 20 μm

Fig. 9

Achillea millefolium L. acute crassus psilate and elongate/oblong/irregular psilate entire/sinuate. This figure demonstrates that these two phytoliths, although broken apart, were once connected while on the plant. Scale bar 50 μm

Tables 1, 2, and 3 list the results of our analysis of phytolith morphotypes produced by each taxon grouped by plant life form, forbs, trees and shrubs and grasses (Utah State University 2017). Following the ICPN 2.0 format (Neumann et al. 2019), all standard morphotype names currently recognized by the ICPN are written in small capitals. In ICPN 2.0 “phytoliths that exhibit features of two closely related morphotypes may bear combined names with descriptors separated by a slash”, so for example, the morphotype Elongate dentate/dendritic would indicate an elongated phytolith with processes that range from dentate or toothed to dendritic or branched; for a discussion of the factors that determine process shape in Elongate, see ICPN 2.0. We also include in each table an initial estimate of the relative abundance or production index (PI) of each phytolith morphotype which was calculated by scanning the sample slides and using a variation of the coding system described by Wallis (2003) and McCune (2013), as follows.

Table 1 Forbs, phytolith morphotypes observed, production indices (PI), and figure references for each species analysed

Table 2 Shrubs and trees, phytolith morphotypes observed, production indices (PI) and figure references

Table 3 Grasses, phytolith morphotypes observed, production indices (PI) and figure references

• Non-producer (NP): no phytoliths observed

• Rare (R): one or two examples of the phytolith morphotype observed on an entire slide

• Uncommon (U): 3–30 of the phytolith morphotype per slide

• Common (C): 30–100 per slide

• Abundant (A): more than 100 per slide

In the ESM, we briefly review the range of plant communities in which each taxon in our study is likely to grow, as well as some of the growth habits and ways in which the taxa were historically used by Great Basin Native Americans.

Forbs

We analysed 14 forbs, non-graminoid herbaceous flowering plants (Table 1). Two, Fragaria vesca L. and Typha latifolia L., were non-producers of phytoliths. The morphotypes we observed most frequently in the other forbs comprising our sample were various epidermal, spheroid ornate and acute trichome phytoliths. None of the morphotypes observed in the forbs were unique to any taxon.

Trees and shrubs

Thirty-one of the plants sampled for this study were trees and shrubs (Table 2). Five species produced no phytoliths. The phytoliths that were produced most frequently were tracheary annulate/helical and spheroid ornate types. A lack of silicification in woody plants has been noted by others (Morris 2008).

Grasses

Seven grass species were analysed (Table 3). Grasses typically produce short-cell and long-cell phytoliths. Five of the grass species that we tested are in the Pooideae subfamily.

Discussion and conclusions

Monocots are known to be the most abundant producers of phytoliths, followed by forbs and woody plants (Pearsall 1989, pp. 360–374). Our findings followed this paradigm. Generally, we found that grasses and forbs were the most common and abundant producers of phytoliths, while shrubs and trees were often non-producers or rare and uncommon producers of phytoliths. Moreover, root and woody samples rarely produced any distinctive phytolith morphotypes. These findings were expected. Accordingly, because some taxa, tissue types, or plant life-forms are underrepresented in the phytolith record, researchers using this reference collection should not attempt to use it to compare the usage of any particular plant life-form to another, or to conduct quantitative analysis. But again, we hope this reference collection will provide a good starting point for any researchers conducting analysis of archaeological phytoliths recovered from Great Basin Native American sites. Such analyses should supplement this reference collection with those of other native wild taxa that grow around the site to assure that similar phytolith morphotypes produced by unused native taxa are not confused for those produced by plants that were used.