Journal of Archaeological Science (2001) 28, 875–886doi:10.1006/jasc.2000.0623, available online at http://www.idealibrary.com on

A Comparative Analysis of Wet and Dry Ashing Techniquesfor the Extraction of Phytoliths from Plant Material

J. F. Parr, C. J. Lentfer & W. E. Boyd*

School of Environmental Science & Management, Southern Cross University, Lismore, New South Wales 2480,Australia

(Received 8 August 1999; revised manuscript accepted 5 October 2000)

Two methods are commonly used for the extraction of phytoliths from plant material to be used as reference in theanalysis of archaeological phytolith samples: (1) spodograms or dry ashings; and (2) acid digestions or wet ashing. Ithas been suggested that these techniques may modify the resultant samples in different ways. Dry ashing, in particular,has been implicated as a cause of shrinkage and warping in phytolith assemblages when incineration occurs at �450�C.The results of a morphometric comparative analysis between the dry ashing and wet ashing methods do not supportthese claims. This study establishes that differences in patterns of dimension and curvature of short bilobate phytolithsand of elongate phytoliths both subjected to dry and wet ash preparation are not statistically significant. There is,therefore, no detectable evidence of morphological impact as a result of these methods. This finding implies that anydifferences that do occur in phytolith size and curvature are typical, possibly random permutation within assemblages,or that they are the result of variation in leaf cell structure rather than the consequence of a particular extractionprocedure. This suggests that the practice of using different methods of preparation of reference samples for fossilanalysis can be reliably continued. � 2001 Academic Press

Keywords: PHYTOLITH ANALYSIS, SAMPLE PREPARATION METHODS, REFERENCE MATERIAL,DRY ASHING, WET ASHING, PHYTOLITH MORPHOLOGY, MORPHOLOGICAL CHANGE.

*Author for correspondence: School of Environmental Science &Management, Southern Cross University, Lismore, New SouthWales 2480, Australia. Tel.: [61-2] 6620 3007; E-mail:bboyd@scu.edu.au

Introduction

V arious methods have been employed for ex-tracting phytoliths from plant material in thepreparation of reference samples for archaeo-

logical fossil phytolith analysis. These may be broadlygrouped into two main categories: the use of spodo-grams or dry ashing; and acid digestion or wet ashing(Royner, 1983: 238; Bowdery, 1989: 172–174). Bothapproaches are well established, and refined variationsof these techniques are currently widely used for thepreparation of phytolith reference collections for plantclassification, palaeoenvironmental reconstruction andarchaeological applications. However, there is somesuggestion that these techniques modify the resultantsamples in different ways (Jones & Milne, 1963: 207–220; Lanning, Hopkins & Loera, 1980: 549–554;Rovner, 1983: 238; Piperno, 1988: 126; Pearsall, 1989:376). This paper reports on the results of a comparativeanalysis between the dry and wet ashing techniques.

Phytolith research has become increasingly moreimportant and widely used in the field of palaeo-environmental analysis (Brown, 1984; Piperno, 1985;

8750305–4403/01/080875+12 $35.00/0

Piperno, Bush & Colinaux, 1991; Wang & Hill,1995; Gol’yeva et al., 1995; Kealhofer & Penny, 1998;Kealhofer & Piperno, 1988), particularly in situationswhere pollen preservation is poor (Boyd, Lentfer &Torrence, 1998). The potential for phytolith analysis tocontribute to the better understanding of prehistoricresource management, food processing and tool func-tion is already evident (Rovner, 1983; Pearsall &Trimble, 1984; Wilson, 1985; Bozarth, 1987; Ball,Brotherson & Gardner, 1993; Piperno & Pearsall,1993; Fullagar, 1993; Rosen, 1994; Pearsall et al., 1995;Kealhofer, 1996; Albert & Weiner, 1997; Owens,1997; Field & Fullagar, 1998; Kealhofer, Torrence& Fullagar, 1999). Moreover, the role of phytolithresearch in plant taxonomy and physiology is wellestablished, and although not as conspicuous perhapsas in other fields of research, it is becoming increasinglypopular (Richardson, 1920; Clifford & Watson, 1977;Ollendorf, Mulholland & Ropp, 1988; Jones & Bryant,1992; Ball, Brotherson & Gardner, 1996).

The integrity of processed samples of fossil phyto-liths is, therefore, fundamental to any research in theareas of palaeoenvironmental reconstruction and ar-chaeological applications and, ultimately, the maincriterion for any phytolith extraction technique mustbe that it will consistently provide morphologically-intact phytoliths which accurately represent original

� 2001 Academic Press

876 J. F. Parr et al.

assemblages (Lentfer & Boyd, 1998: 1159). Neverthe-less, potential differences in the preparation of phyto-liths for analysis have been suggested to occur betweenextraction methods. In particular, the dry ashing tech-nique has been implicated as a cause of shrinkage,warping and changes to the refractive index of phyto-liths (Jones & Milne, 1963: 207–220; Jones &Handreck, 1967: 125–126; Pearsall, 1979: 146–147,1989: 375). Such an effect of the extraction methodmay seriously impede identification and accurateanalysis of fossil phytoliths. Although this hypothesishas been questioned (Lanning, Hopkins & Loera,1980: 550; Piperno, 1988: 126–127; Runge, pers.comm.), there has been no systematic study carried outto test the assumption of these effects.

Dry ashing was the original method employed toseparate phytoliths from their surrounding organicmatter. This technique has evolved from Mohl’s (1861)experimentation on the retrieval of phytoliths from theremnant ash of burnt plant material. Subsequent re-searchers, such as Schelenberg (1908), Molisch (1920),Richardson (1920) and Policard (1923), examined arange of dry ashing techniques (described by Uber,1940: 204, and Netolitsky, 1929; both cited by Jones &Handreck, 1967: 125; Piperno, 1988: 126; Bowdery,1989: 172). The procedures that are fundamental to dryashing include the pre-treatment and/or post-treatments of samples by washing with distilled waterand/or hydrochloric acid (HCl); additional productshave also been used such as hydrogen peroxide (H2O2)(see Bowdery, 1989: 174). Samples are placed in cruci-bles, in a muffle furnace and ignited at a desiredtemperature. Incineration rates are usually maintainedat about 500�C (Stewart & Arthur, 1935: 905; Jones &Handreck, 1967: 125; Piperno, 1988: 126; Pearsall,1989: 377). Nevertheless, Rovner (1983: 238) pointsout that higher temperatures up to 1000�C have beenapplied; an example is provided by Lanning, Ponnaiya& Crumpton (1958: 339) who have used temperaturesin the range of 700–900�C.

The concept of wet ashing was introduced early thiscentury by Zimmerman (1901), although it was notwidely recognized until the late 1950s (e.g. Parry &Smithson, 1957: 976) and increased in popularity withthe work of Jones & Milne (1963) and Rovner (1972).Wet ashing also requires the pre- and/or post-treatments described above, while the incinerationstage is substituted by the use of acid digestion as ameans of oxidation (e.g. Jones & Milne, 1963: 209).Bowdery (1989: 174–175) provides tables outlining theranges of temperatures, times and chemicals used inthese techniques and their associated references.

Over the last four decades there has been ongoingdebate regarding the advantages and disadvantagesof these techniques (Jones & Milne, 1963: 207–220;Lanning, Hopkins & Loera, 1980: 549–554; Rovner,1983: 238; Piperno, 1988: 126; Pearsall, 1989: 376).Piperno (1988: 126–127), for example, has questonedthe need for concern in relation to the dry ashing

process, suggesting that previous research (e.g.Lanning, Hopkins & Loera, 1980: 550) demonstratedthat if firing temperatures do not exceed 500�C, mor-phological changes in phytoliths do not occur. A recentstudy testing a range of temperatures on phytolithsfrom African arboreal species supports this proposi-tion (Runge, E., pers. comm.). In contrast to the dryashing debate, some researchers have found difficultiesin applying the wet ashing technique, in particular,commenting on difficulties in the use of Schulze solu-tion (a combination of nitric acid (HNO3) and potass-ium chlorate (KClO3) or sodium chlorate (NaClO3);(Pearsall, 1989: 381; McWeeney, pers. comm.). One ofthe authors (Lentfer) has also encountered compli-cations with the use of sodium hypochlorite (bleach)for digestion. The problematic areas associated withwet oxidation appears to revolve around differentialdigestion rates of phytoliths, cellulose, and other planttissue by the chemicals used for wet ashing, andapparent loss of phytolith residue (Pearsall, 1989: 381).Rosen (pers. comm.) has found that dry ashingyields around 20% more phytoliths than wet ashing.Finally, a further area of concern for some phytolithresearchers is the use of noxious chemicals necessaryfor the wet ashing process (Piperno, 1988: 126; Runge,pers. comm.).

It is clear, however, that both techniques have theirstrengths and weaknesses. Their application will varydepending on the nature of the research question andthe personal preferences of the analyst. Rosen (pers.comm.), for example, has suggested that dry ashingmay be more applicable for comparative referencematerial in relaiton to archaeological sites becausemany of their phytoliths are retrieved from ash de-posits. Alternatively, Rovner (pers. comm.) points outthat not all phytoliths recovers from archaeologicaldeposits have been subjected to fire, and in these caseswet ashing is a more appropriate method.

Following research initiated by Lentfer (1995), thispaper examines both ashing techniques in order toassess the nature of evidence for phytolith modifi-cation, especially in regard to shrinkage and warping.A systematic comparative analysis of phytolith mor-phology after dry and wet ash extractions will assist inthe understanding of their individual capacities, andmost importantly, the ability of these extraction tech-niques to produce comparative results. It is consideredessential amongst phytolith analysts that this issue beaddressed to clarify any ambiguity that may affectphytolith analyses (Hodson, Rovner & Rosen, pers.comm.).

MethodsBoth extraction methods applied in this assessment arebased on frequently-used methodologies with the aimof allowing replication using standardized proceduresfor phytolith analysis. Leaf samples were cut up

Comparative Analysis of Wet and Dry Ashing 877

Table 1. Description of dry and wet ash phytolith extraction from plant material

Dry ashing Wet ashing

Extraction method using fume hood burning with distilled water,10% HCl and 15% H2O2 for washing, the removal of carbonates,and organics respectively.

Extraction method using Schulze solution i.e. nitric acid (HNO3)and potassium chlorate (KClO3). Washing with distilled water.

(1) Weigh test tube and record.(2) Weigh dried plant material and record.(3) Rinse plant material in distilled water and transfer to

crucible.(4) Make template for fume hood.(5) Transfer lidded crucibles to fume hood.(6) Heat to 500�C and hold for 6 h.(7) Switch off fume hood and leave overnight to cool.(8) Remove crucibles and transfer contents to test tubes.(9) Add 10 ml of 10% HCl to test tube—heat in water

bath@70�C for 20 min or until reaction stops.(10) Centrifuge@3500 rpm for 5 min and decant.(11) Rinse with distilled water and centrifuge@3500 rpm for

5 min and decant.(12) Add 10 ml of 15% H2O2—heat in water bath@70�C for

20 min or until reaction stops.(13) Centrifuge@3500 rpm for 5 min and decant.(14) Rinse with 10 ml of distilled water and centrifuge@3500 rpm

for 5 min and decant. Repeat rinse.(15) Add 1 ml of ETOH and leave overnight to dry.(16) Weigh dried material, calculate phytolith weight and transfer

to labelled vials with as little 100% ETOH as possible.

(1) Weigh test tube and record.(2) Weigh dried plant material and record.(3) Rinse plant material in distilled water and transfer to flask.(4) Add 10 mls of HNO3 and a pinch of KClO3.(5) Place on hotplate set to simmering temperature. Stir

regularly—when contents changes from its initial brown ororange state to clear yellow transfer contents to test tube.

(6) Centrifuge@3500 rpm for 5 min and decant.(7) Rinse with 10 ml of distilled water and centrifuge@3500 rpm

for 5 min and decant. Repeat rinse two times.(8) Add 1 ml of ETOH and leave overnight to dry.(9) Weigh dried material, calculate phytolith weight and transfer

to labelled vials with as little 100% ETOH as possible.

finely, mixed to ensure homogeneity, and then dividedinto equal weight portions so that both dry and wetoxidation extraction techniques could be applied. Adescription of the extraction methods is provided inTable 1. The dry ashing technique used in this studywas a slightly modified version of that used by Twiss,Suess & Smith (1969: 111), with the addition of centri-fuging, hydrochloric acid (HCl) and hydrogen peroxide(H2O2) after initial ashing (Hart, 1998: 9). The reasonfor the additional use of HCl and H2O2 was to provideclean phytolith assemblages and to assist lucidity dur-ing morphometric analysis. Wet ashing was achievedthrough the use of a method similar to that ofRovner (1972: 591), with Schulze solution (nitric acidand potassium chlorate) employed for digestion(Table 1).

The plant materials used in this study were obtainedfrom the Herbarium at the Lae Forestry ResearchInstitute, Morobe Province, Papua New Guinea. Allsamples are voucher specimens from that Institute,with species identification and accession numbers pro-vided; only very small amounts of plant materials wereavailable for samples (Table 2). The specimens com-prise the leaf sections of 10 different Poaceae species.However, due to the lack of suitable phytolith types insome species, from these 10 species only eight wereselected for each of the two analyses, shrinkage andwarping. The selection criterion for species in eachstudy was that they comprise sufficient quantities ofshort and long celled phytoliths respectively.

To test the assumption that dry ashing and wetashing extraction techniques produced comprative re-sults, two separate analyses were conducted assessingpatterns of morphometric change following process-ing. The first analysis compared the size of bilobate(dumbbell) shaped phytoliths (Twiss, Suess & Smith,1969: 111), and the second measured the curvature ofelongate phytoliths after extraction procedures. Eightspecies were chosen for each analysis according toyields of short cells and long cells. Only those specieswith high yields were suitable for study. Phytolithresidue assemblages were mounted on to glass slides inEukitt mounting medium. Slides were viewed at 400�magnification with an Olympus CH-2 compound mi-croscope fitted with a polarizing filter and a cameralucida. From each slide, one area containing phytolithswas photographed for later reference using anOlympus camera mounted to a photo tube on anOlympus BH.2 microscope at 400� magnification.Phytoliths were then traced in plain view using thecamera lucida, and divided into two categories (i.e.short cells and long cells). The overall sample com-prised 1600 phytoliths, with 50 from each of the twoextraction methods analysed for each species. Twoseparate analyses were conducted to assess the natureof phytolith morphology following dry and wet ashingextraction methods. The aim was to test the assump-tion that both dry and wet ashing techniques producedcomparative results. The first analysis compared thesize of bilobate (dumbbell) phytoliths (Twiss, Suess

878 J. F. Parr et al.

& Smith, 1969: 111), and the second measured thecurvature of elongate phytoliths after extractionprocedures.

Width and length comparisonsBilobate phytoliths were selected to represent the shortcell category because they were present in sufficientquantities for statistical analysis in eight of the 10species assessed. The short cells were designated to testfor shrinkage only, because they are compact formsand less likely to warp. Measurements were taken ofthe length (i.e. maximum dimension (MD) and thewidths across the two widest points at each end of thephytolith. The two width measurements were reducedto mean width (MMW) for the analysis (Figure 1).

Comparison of mean anglesLong cells, mostly spiny elongates (Twiss, Suess &Smith, 1969: 111), were selected to test for warping. It

was anticipated that warping would be more pro-nounced in elongate phytoliths and thus that theywould provide a clearer evidence for distortion. Theelongate phytoliths were traced using a camera lucida,digitized using a Sharp JX-320 flatbed scanner andthen imported into the program Image Tool on aPower Macintosh PC. Image Tool parameters were setto measure the degree of curvature. Calibrations weremade for angles at each end of elongate phytoliths(Figure 2).

Table 2. Summary of Poaceae species, authority, accession numbers Herbarium Lae Forestry Research Institute, type, fresh weight, residue weightand extraction method

Species Accession no. Sample type Sample wt Residue wt Method

Bambusa sp. LH. 103886 Leaf 0·1000 g 0·0065 g DALeaf 0·1000 g 0·0107 g WA

Brachiaria brizantha (Hoscht. ex A. Rich) Stapf. LH. 242415 Leaf 0·0610 g 0·0036 g DALeaf 0·0610 g 0·061 g WA

Buergersiochloa macrophylla S.T. Blake, Blumea Supp. LH. 229126 Leaf 0·1000 g 0·0037 g DALeaf 0·1000 g 0·0112 g WA

Heteropogon triticus (R.Br) Stapf. ex Craib LH. 18775 Leaf 0·1000 g 0·0018 g DALeaf 0·1000 g 0·0145 g WA

Imperata exaltata (Roxb.) Brogn. LH. 81921 Leaf 0·1000 g 0·0024 g DALeaf 0·1000 g 0·0213 g WA

Polytoca macrophylla Benth. LH. 243442 Leaf 0·0768 g 0·0088 g DALeaf 0·0768 g 0·0113 g WA

Saccharum robustum LH. 18598 Leaf 0·0772 g 0·0087 g DALeaf 0·0772 g 0·003 g WA

Saccharum officinarum (L.) LH. 94173 Leaf 1·000 g 0·0001 g DALeaf 1·000 g 0·011 g WA

Themeda arguens (L.) Hack. LH. 12057 Leaf 0·0267 g 0·0012 g DALeaf 0·0267 g 0·0013 g WA

Thysanoleana maxima (Roxb.) O.K. LH. 207698 Leaf 0·1000 g 0·0055 g DALeaf 0·1000 g 0·0177 g WA

Figure 1. Illustration of parameters measured for the size analysis ofbilobate phytoliths.

Figure 2. Illustration of parameters measured for the angle ofdistortion analysis of elongate phytoliths using the Image Toolsoftware.

Statistical analysisStatistical analyses are based on a comparative mor-phology approach similar to that used by Lepofsky,Kirch & Lertzman (1998: 1006–1007). A two-tailedt-test using SPSS for statistical analysis compares themean dimensions of bilobates, the mean angles of

Comparative Analysis of Wet and Dry Ashing 879

elongate phytoliths, and boxplots are used to providevisual comparisons of the mean maximum width(MMW), maximum dimension (MD) length, and meanangle distribution after dry and wet ashing procedures.Each box defines the interquartile range i.e. iqr=upperquartile plus lower quartiles. The whiskers extendingfrom above and below the iqr represent the extent ofthe 100th and 0 percentiles respectively. Outlyingvalues denoted by diamonds and asterisks representvalues that all fall further than 1·5 times and 3 timesthe interquartile range from the end of the box respect-ively (Devore & Peck, 1993: 97–99).

Figure 3. Examples of a dry ashed sample, showing clear sheets ofelongate phytoliths with in-situ bilobates and stomates.

Figure 4. Example of a dry ashed sample, showing disarticulatedshort cells, including bilobates and crosses.

Figure 5. Example of a wet ashed sample, showing aggregated sheetsof unclear elongate phytoliths with in-situ bilobates.

Figure 6. Example of a wet ashed sample, showing disarticulatedshort cells, including bilobates and crosses.

ResultsProcessing times based on actual duration spent work-ing in the laboratory was comparatively similar fordry and for wet ashing, i.e. around 3 h. The datapresented in Table 2 shows that all the correspondingsamples started with the same plant material weight.After phytolith extraction the dry ashing methodresulted in consistently lower residue weight than thewet ashing. The wet ash residues had a mean averageweight of 63·7% greater than those for dry ashedsamples.

Microscopic analysis at 100� and 400� magnifi-cations found that in the dry ashed samples, elongatephytoliths were mainly in sheets with in-situ bilobates,stomates, and bulliforms, as well as disarticulated cellssuch as cross shapes, tribolates, and trichomes (Figures3 & 4). Most phytoliths in the dry ashed samples weresufficiently lucid and free of residual material. The useof polarized light at 400� and 100� magnificationsfailed to detect calcium oxalate crystals or starch grainsin these samples. Wet ashed samples had comparabledisarticulated elongates and short cells, but in-situphytoliths mainly occurred in large, relatively un-diagnostic, amber-coloured clumps (Figures 5 & 6).Under crossed polarized light at 400� magnification,these clumps comprised elongate phytoliths bound inwhite matted material. At 100� magnification the

white matted areas were shown to consist of individualelongated strips, not raphides, but cellulose-likematerial that apparently had not been oxidized. Occa-sional raphides and starch grains were observed ran-domly throughout wet ashed samples. During themicroscopic analysis it was also noted that manyelongate phytoliths in both wet and dry ashed sampleshad a large degree of curvature when in associationwith stomates.

880 J. F. Parr et al.

Width and length comparisons for shrinkageThe results of the two-tailed t-tests and boxplotscomparing dry ashing and wet ashing and testing forshrinkage are shown in Table 3 and Figure 7. Differ-ences in the MMW for Heteropogon riticus, Imperataexaltata, Saccharum robustum, Themeda arguens, andSaccharum officinarum samples were significant atP=0·05 (Table 3). The dry ashed MMWs are signifi-cantly larger for H. triticus and S. robustum (P=0·000),yet smaller for I. exaltata (P=0·005), T. arguens(P=0·000) and S. officinarum (P=0·018). No signifi-cant differences occur in MMW between the wet anddry ashed samples of Buergersiochloa macrophylla(P=0·100), Brachiaria brizantha (P=0·887) and Thysa-noleana maxima (P=0·670). The boxplots show thatdry ashed samples of H. triticus, B. brizantha, B.macrophylla and T. maxima have significantly largewidth distributions in comparison to wet ashedsamples. Alternatively, I. exaltata, S. robustum andT. arguens have comparatively small width distribu-tions (Figure 7). Results for the MD (Table 3) showthat the dry ashed samples are significantly larger forH. triticus (P=0·000) and S. officinarum (P=0·009),but smaller for I. exaltata (P=0·048) and S. robustum(P=0·000). There are no significant differencesbetween methods for T. arguens (P=0·404), B. macro-phylla (P=0·564), B. brizantha (P=0·652) and T.maxima (P=0·690). The boxplots demonstrate thatthe MD distributions for the dry ashed samplesof H. triticus, B. brizantha, S. robustum, S. officinarumand T. arguens are large in comparison to wetashed samples. The dry ashed samples of B. macro-phylla, I. exaltata, and T. maxima have compara-tively small distributions to wet ashed samples(Figure 7).

Comparison of mean angles for warpingThe results of the two-tailed t-tests and boxplotscomparing dry ashing and wet ashing and testing forwarping are shown in Table 4 and Figure 8. Statisticalanalysis found that the angle of curvature for three ofthe eight species examined had significantly differentP-values (Table 4). Mean angles were significantlylarger for the dry ashed sample of H. triticus(P=0·020), but smaller for P. macrophylla (P=0·002)and S. officinarum (P=0·044). No significant differ-ences occur in the results for Bambusa sp. (P=0·051), I.exaltata (P=0·110), S. robustum (P=0·239), T. arguens(P=0·111) and T. maxima (P=0·378). The boxplotsshow that, if the few outliers that occur are discounted,the curvature and angle distribution is less for dryashed specimens in five of the eight species assessed(Figure 8). These five species comprise I. exaltata,P. macrophylla, S. robustum, S. officinarum andT. maxima. Of the remaining species, T. arguens andBambusa sp. are comparatively proportional, whileH. triticus has a relatively larger angle distribution thanthe wet ashed samples.

Table 3. Summary statistics of analysis testing degree of shrinkage in bilobate phytoliths for dry ashing and wet ashingextraction methods

Species Method

Mean maximum width (�m) Maximum dimensions (�m)

x̄�S.D. Range N Sig. x̄�S.D. Range N Sig.

Heteropogon triticus DA 10·8�2·0 7·5–15·0 50 0·000 18·1�2·2 12·5–22·5 50 0·000WA 9·4�1·4 9·8–12·5 50 16·2�2·7 11·2–22·5 50

Brachiaria brizantha DA 12·6�2·0 7·5–17·5 50 0·887 19·2�2·8 12·5–25·0 50 0·652WA 12·6�1·4 10·0–17·5 50 19·4�2·7 12·5–25·0 50

Buergersiochloa macrophylla DA 13·4�3·8 6·2–25·0 50 0·100 22·0�5·0 10·0–30·0 50 0·564WA 14·6�3·4 7·5–22·5 50 21·4�4·3 15·0–31·2 50

Imperata exaltata DA 9·5�1·5 7·5–13·7 50 0·005 15·6�2·7 11·2–22·5 50 0·048WA 10·6�2·1 6·2–16·2 50 16·7�12·5 12·5–23·7 50

Saccharum robustum DA 11·2�1·4 8·1–15·0 50 0·000 28·1�3·6 20·3–37·5 50 0·000WA 13·2�2·1 8·1–18·7 50 20·8�2·6 16·2–27·5 50

Saccharum officinarum DA 14·7�2·1 10·0–20·0 50 0·018 27·7�4·8 20·0–37·5 50 0·009WA 15·7�2·0 11·2–20·0 50 25·4�3·7 16·2–33·7 50

Themeda arguens DA 9·3�1·2 6·2–13·3 50 0·000 20·8�3·6 15·0–31·2 50 0·404WA 11·4�1·7 6·2–15·0 50 21·4�3·7 15·0–30·0 50

Thysanoleana maxima DA 11·5�2·2 6·2–17·5 50 0·67 19·2�2·3 12·5–23·7 50 0·69WA 11·7�2·0 7·5–16·2 50 19·4�2·8 12·5–25·0 50

DiscussionResidue quantities

Residue weights for the wet ashed samples are consist-ently larger than those for dry ashing (Table 2), a resultwhich is in contradiction to results reported by Rosen(pers. comm.), who found that dry ashing providedaround a 20% increase in phytolith retrieval whencompared to wet ashing. Here wet ashing has providedan average of 63·7% more residue weight than dryashing. Hodson (pers. comm.) has suggested that there

Comparative Analysis of Wet and Dry Ashing 881

0

40

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Buergersiochloa macrophylla

Wet AshMD

Wet AshMMW

Dry AshMD

Method

30

0

30

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Heteropogon triticus

Wet AshMD

Wet AshMMW

Dry AshMD

Method

0

40

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Saccharum robustum

Wet AshMD

Wet AshMMW

Dry AshMD

Method

30

0

40

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Themeda arguens

Wet AshMD

Wet AshMMW

Dry AshMD

Method

30

0

30

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Imperata exaltata

Wet AshMD

Wet AshMMW

Dry AshMD

Method

0

40

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Saccharum officinarum

Wet AshMD

Wet AshMMW

Dry AshMD

Method

30

0

30

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Thysanolaena maxima

Wet AshMD

Wet AshMMW

Dry AshMD

Method

0

30

Len

gth

(m

icro

ns)

Dry AshMMW

10

20

Brachiara brizantha

Wet AshMD

Wet AshMMW

Dry AshMD

Method

Figure 7. Boxplots illustrating sample distributions according to mean maximum widths of bilobates (MMW) and maximum dimensions (MD)for dry ashing and wet ashing. The central box showd the median and the lower and upper quartiles of the distribution. Outlying values shownby diamonds and asterisks represent values that fall further than 1·5 times and 3 times the interquartile range from the ends of the boxrespectively.

882 J. F. Parr et al.

is a greater possibility of loss in the number of lightlysilicified cell wall deposits as a result of acid digestion.The higher yield of compound plates with dry ashingprocedures in comparison to wet ashing (Raeside,1970: 125) supports Hodson’s view. However, an im-portant question follows: if it is assumed that dryashing produces larger phytolith yields, why are theresidue weights in this study consistently greater in thewet ashed samples? The white matted areas ofcellulose-like material mentioned in the results onlyoccurred in the wet ashed samples. The failure of thismaterial to oxidize using the Schulze solution mayaccount for the higher residue weight for wet ashedsamples. The ability of some plants to withstand thewet oxidation process has previously been recognizedas a problematic area (Pearsall, 1989: 381). It is poss-ible that the wet ashing method may have needed to beapplied for a longer duration and/or additional KClO3applied to promote greater reaction during oxidaton.This implies that this stage of the wet ashing processrequires a combination of regular monitoring andchemical adjustment to suit individual samples. On theother hand dry ashing produces consistent resultsaccording to a standardized set of procedures, and istherefore the simpler method of the two.

Table 4. Summary statistics of analysis testing degrees of curvature in elongate phytoliths for dry ashing and wetashing extraction methods

Species

Mean curvature

N Sig.Method x̄�S.D. Range

Bambusa sp. DA 3·9�4·2 0·01–16·8 50 0·051WA 5·8�4·9 0·01–18·2 50

Heteropogon triticus DA 5·0�4·2 0·00–15·9 50 0·02WA 3·2�3·3 0·05–11·5 50

Imperata exaltata DA 4·5�2·8 0·09–12·08 50 0·11WA 5·8�4·8 0·09–18·9 50

Polytoca macrophylla DA 3·0�3·4 0·07–15·2 50 0·002WA 5·8�4·9 0·03–22·8 50

Saccharum robustum DA 4·5�5·3 0·02–27·7 50 0·239WA 5·7�5·1 0·04–19·1 50

Saccharum officinarum DA 4·2�4·6 0·05–21·3 50 0·044WA 6·2�5·3 0·00–19·2 50

Themeda arguens DA 3·0�3·0 0·04–11·7 50 0·111WA 4·3�4·5 0·00–18·28 50

Thysanoleana maxima DA 4·46�4·0 0·02–18·05 50 0·378WA 5·2�4·9 0·02–18·5 50

ShrinkageUnlike the results for residue weights, there was noevidence that either dry ashed or wet ashed samplesconsistently produced larger or smaller MMW or MDvalues. Five of the eight species assessed in this analysisproduced significantly different results according to thetreatments. Nevertheless, these results appear to berandom rather than the consequence of a particularextraction procedure. For example, statistical analysis

for MMW indicated that dry ashed samples are sig-nificantly larger for Heteropogon triticus, smaller forImperata exaltata, larger for Saccharum robustum,smaller for Themeda arguens, and smaller for Saccha-rum officinarum. The analysis of MD resulted in asimilarly random pattern: for dry ashing, samples forH. triticus and S. officinarum are larger, I. exaltata andS. robustum are smaller, with there being no significantdifference between methods for T. arguens.

Although comparison between dry and wet ashingresults for shrinkage reveal significant differences bothbetween individual samples and groups of assemblages,the lackof recurring patterns in the results from eithermethod indicates that both differences in width andlength probably do not represent size modification dur-ing extraction procedures. The inconsistent pattern ofvariation, therefore, most likely reflects the naturalrange of variability in the phytolith residues them-selves, rather than modification due to the applicationof the processing treatments. Although the samplesfrom each species were cut up finely and were acquiredfrom the same portion of leaf, it is relevant to note thatthe leaf segments have a range of variation across theircell structure and that some species have notable vari-ability in cell morphology. Whang, Kim & Hess (1998:465) have found, for example, that in the leaves of somePoaceae species there are significant differences in phy-tolith morphology depending on their proximity tomidribs or lateral veins. Similar results in other speciesare reported by Ball, Brotherson & Gardner (1993),Ball, Gardner & Brotherson (1996) and Whang & Hill(1995). Thus it is possible that the results shown hereare an artefact of such variation and occurred despiteefforts to ensure homogeneity of samples prior totreatment during their preparation. This emphasizes,therefore, the importance of selection of plant material

Comparative Analysis of Wet and Dry Ashing 883

–10

30

An

gle

of c

urv

atu

re

10

20

Thysanolaena maxima

Method

0

Dry Ash Wet Ash–10

20

An

gle

of c

urv

atu

re

0

10

Themeda arguens

MethodDry Ash Wet Ash

–10

30

An

gle

of c

urv

atu

re

0

10

Saccharum robustum

Method

20

Dry Ash Wet Ash–10

30

An

gle

of c

urv

atu

re

0

10

Saccharum officinarum

Method

20

Dry Ash Wet Ash

–10

30

An

gle

of c

urv

atu

re

0

Imperata exaltata

Method

10

20

Dry Ash Wet Ash–10

30

An

gle

of c

urv

atu

re

0

10

Polytoca macrophylla

Method

20

Dry Ash Wet Ash

–10

20

An

gle

of c

urv

atu

re

Dry Ash

0

10

Heteropogon triticus

Wet AshMethod

–10

20

An

gle

of c

urv

atu

re

0

10

Bambusa sp.

MethodDry Ash Wet Ash

Figure 8. Boxplots illustrating sample distributions according to degree of curvature measured in elongate phytoliths. The layout is asfor Figure 7.

884 J. F. Parr et al.

for reference collections, and the need for carefullyplanned, systematic sampling to ensure the full range ofmorphological types are accounted for.

WarpingThis analysis assessed the elongate phytoliths for evi-dence of warping as a result of preparation technique.The computer-assisted image analysis was a particu-larly efficient way to measure microscopic featuressuch as the curvature of phytoliths and transfer thisdata directly into statistical packages. Statistical testingfound that only three of the eight species examinedwere significantly different according to the treatments.The statistical analysis for these three species showthat the degree of curvature of phytoliths in the dryashed sample was significantly greater for H. triticussignificantly greater for H. triticus, but lesser forP. macrophylla and S. officinarum. Again this appearsto be random patterning similar to that observed inthe previous analysis. However, comparisons madebetween dry and wet ashed samples using the boxplotsin Figure 8, indicate that with the exception of tworelatively extreme outliers in S. robustum and S. offici-narum, only one of the eight dry ashed species assessedin this analysis, H. triticus, had a significantly largerdistribution and degree of curvature. Dry ashedsamples Bambusa sp. and T. arguens had no significantdifference to their corresponding wet ashed boxplots.The remaining five species had both a significantlylarger distribution of curved elongates and exhibited alarger degree of individual curvature in wet ashedsamples. Again this may be the result of variation inleaf cell structure (Ball, Brotherson & Gardner, 1993;Ball, Gardner & Brotherson, 1996; Whang & Hill,1995); plates within a number of publications show,for example, curved elongate phytoliths in-situ withstomates and other features from both dry and wetashed samples (e.g. Jones & Handreck, 1967: 128;Clifford & Watson, 1977: 45–77; Lanning, Hopkins &Loera, 1980: 554). The results, therefore, from theseanalyses do not support the claim that dry ashingresults in shrinkage and warping of phytoliths ataround �500�C as proposed by Jones & Milne (1963:207–220). As suggested by Lanning, Hopkins & Loera(1980: 550), Piperno (1988: 126–127) and Runge (pers.comm.), there appears to be little need for concern inrelation to the dry ashing process, if firing temperaturesdo not exceed 500�C.

ConclusionThis comparison of the results for wet and dry ashingprocedures has shown that the dry ashing methodproduces less residual matter than the wet ashingtechnique. Moreover, it provides clean, lucid, disarticu-lated, and in-situ phytolith assemblages more effec-tively than does the wet ashing method. This outcome

is highly desirable for comprehensive reference collec-tions, and in particular for those which will be used toprovide detailed information on phytolith location andvariation within plants. Both methods are comparablein time taken to carry out the phytolith extractionprocedures, and importantly, it is indicated here thatwet ashing also produced comparable results to thoseof dry ashing for disarticulated phytoliths.

However, wet ashed samples do not allow the in-situmaterial to be clearly distinguishable due to the aggre-gation of residual matter that apparently had not beenfully oxidized. This implies that a combination ofregular monitoring and chemical adjustment to suitindividual samples during preparation is necessary toachieve satisfactory results with the wet ashing process.In contrast, the dry ashing treatment produces consist-ent results and can be standardized for all plants andplant parts.

This study has, importantly, established that differ-ences in patterns of dimension and curvature of shortbilobate phytoliths and longer elongate phytoliths sub-jected to dry and wet ash preparation are not statisti-cally significant. There is no detectable evidence fromthis study, therefore, of morphological impact as aresult of the application of methods. This implies thatany differences that do occur in phytolith size andcurvature may be considered to be typical, possiblyrandom permutations within the original assemblagesof phytoliths, in part probably assigned to variation inleaf cell structure rather than as a consequence of aparticular extraction procedure. Such conclusionsimply, therefore, that while care may be needed in theselection of sample preparation method in terms ofthe eventual use of the reference sample, in terms ofproviding morphologically comparable referencematerial, the practice of using various methods ofpreparation of reference samples for fossil analysis canbe reliably continued, and that the concerns expressedby various practitioners may be allayed.

AcknowledgementsThe authors also wish to acknowledge with gratitude,the contributions to this discussion of members of thee-mail discussion list PHY-TALK, who promptly com-mented to an initial question regarding the topic of thispaper transmitted over this e-mail list. In particular,the following people are thanked for their specificcontributions: Freya Runge, Lucinda McWeeney,Arlene Rosen, Iv Rovner and Martin Hodson. TheHerbarium at the Lae Forestry Research Institute,Morobe Province, Papua New Guinea made an invalu-able contribution to this work in its supply of her-barium plant specimens. This project was in partfunded from an ARC Large Grant awarded to Boyd tosupport the ‘‘Prehistory of Garua, Papua New GuineaArchaeological Project’’ of which the research reportedhere is but a small part.

Comparative Analysis of Wet and Dry Ashing 885

ReferencesAlbert, R. M. & Weiner, S. (1997). Phytoliths in wood ash. The

Phytolitharien 10, 5–6.Clifford, H. T. & Watson, L. (1977). Identifying Grasses: Data,

Methods and Illustrations. Brisbane: University of QueenslandPress.

Ball, T. B., Brotherson, J. D. & Gardner, J. S. (1993). A typologicand morphometric study of variation in phytoliths from einkornwheat (Triticum monococcum). Canadian Journal of Botany 71,1182–1192.

Ball, T. B., Gardner, J. S. & Brotherson, J. D. (1996). Identifyingphytoliths produced by the inflorescence bracts of three speciesof Wheat (Triticum monococcum L., T. dodoccon Schrank., andT. aestivum L.) using computer-assisted image and statisticalanalyses. Journal of Archaeological Science 23, 619–632.

Bowdery, D. (1989). Phytolith analysis: introduction and applica-tions. In (W. Beck, A. Clarke & L. Head, Eds) Plants in AustralianArchaeology. Brisbane: Watson Ferguson & Company, pp. 161–186.

Boyd, W. E., Lentfer, C. J. & Torrence, R. (1998). Phytolith analysisfor a wet tropics environment: methodological issues and implica-tions for the archaeology of Garua Island, West New Britain,Papua New Guinea. Palynology 22, 213–228.

Bozarth, S. R. (1987). Diagnostic opal phytoliths from rinds ofselected Cucurbita species. American Antiquity 52, 607–615.

Brown, D. A. (1984). Prospects and limits of a phytolith key forgrasses in the central United States. Journal of ArchaeologicalScience 11, 345–368.

Devore, J. & Peck, R. (1993). Statistics: The Exploration andAnalysis of Data, 2nd edn. Belmont: CaliforniaDuxbury Press.

Field, J. & Fullagar, R. (1998). Grinding and pounding stones fromCuddie Springs and Jinmium. In (R. Fullagar, Ed.) A Closer Look:Recent Australian Studies of Stone Tools. Sydney UniversityArchaeological Methods Series 6. Australia: University of Sydney.

Fullagar, R. (1993). Taphonomy and tool-use: a role for phytolithsin use-wear and residues analysis. In (B. L. Frankhauser &S. R. Bird, Eds) Archaeometry: Current Australasian Research.Occasional Papers in Prehistory No. 22, pp. 21–27.

Gol’yeva, A. A., Aleksandrovskiy, A. L. & Tselishcheva, L. K.(1995). Phytolithic analysis of holocene paleosoils. Eurasian SoilScience 27, 46–57.

Hart, D. (1998). Sample preparation techniques. In (D. Hart & G. S.Humphreys, Eds) Australiasian Phytolith Workshop, Feb. 9 & 10,1998. Sydney: Geoecology Group, School of Earth Sciences,Macquarie University.

Jones, J. G. & Bryant, V. M. Jr (1992). Phytolith taxonomy inselected species of Texas cacti. In (G. Rapp Jr & S. C. Mulholland,Eds) Phytolith Systematics. New York: Plenum Press.

Jones, L. & Handreck, K. (1967). Silica in soils, plants and animals.Advances in Agronomy 19, 107–149.

Jones, L. H. P. & Milne, A. A. (1963). Studies of silica in the oatplant. Plant and Soil XVIII, 207–220.

Kealhofer, L. (1996). The human environment during the terminalpleistocene and holocene in northeastern Thailand: phytolithevidence from Lake Kumphawapi. Asian Perspectives 35, 229–254.

Kealhofer, L. & Penny, D. (1998). A combined pollen and phytolithrecord for fourteen thousand years of vegetation change in north-eastern Thailand. Review of Palaeobotany and Palynology 103,83–93.

Kealhofer, L. & Piperno, D. R. (1988). Opal Phytoliths in Southeast-ern Asian Flora. Smithsonian Contributions to Botany. No. 88.

Kealhofer, L., Torrence, R. & Fullagar, R. (1999). Integratingphytoliths within use-wear/residue studies of stone tools. Journalof Archaeological Science 26, 527–546.

Lanning, F. C., Hopkins, T. L. & Loera, J. C. (1980). Silica and ashcontent and depositional patterns in tissues of mature Zea mays L.plants. Annals of Botany 45, 549–554.

Lanning, F. C., Ponnaiya, B. W. X. & Crumpton, C. F. (1958). Thechemical nature of silica in plants. Plant Physiology 33, 339–343.

Lentfer, C. J. (1995). Ethnobotanical research in West New Britainand the Forestry Research Institute Herbarium, Lae, PNG. Unpub-lished Report. Sydney, NSW: Australian Museum.

Lentfer, C. J. & Boyd, W. E. (1998). A comparison of three methodsfor the extraction of phytoliths from sediments. Journal ofArchaeological Science 25, 1159–1183.

Lepofsky, D., Kirch, P. V. & Lertzman, K. P. (1998). Metricanalyses of prehistoric morphological change in cultivated fruitsand nuts: an example from Island Melanesia. Journal of Archaeo-logical Science 25, 1001–1014.

Mohl, H. von. (1861). Uber das Kieselskelett lebender pflanzenzel-len. Botanische Zeitung 19, 209–215.

Molisch, H. (1920). Aschenbild und pflanz enverwandtschaft. Sit-zungsberichte Akademie der Wissenschaften 129, 261–294.

Netolitsky, F. (1929). Die Kieselkorper die Kalksalze als Zellinhald-skorper. In (K. Linsbauer, Ed.) Handbuch der Pflanzenanatomie,3.. Berlin: Gebruder Borntraeger, pp. 1–19.

Ollendorf, A. L., Mulholland, S. C. & Rapp, G. Jr (1988). Phytolithanalysis as a means of plant identification: Arundo donax andPhragmites communis. Annals of Botany 61, 209–214.

Owens, D. (1997). A feasibility study for phytolith research in theSoutheast from Scull Shoals in the Oconee National Forest,Georgia. The Phytolitharien 10, 6–8.

Parry, D. W. & Smithson, F. (1957). Detection of opaline silica ingrass leaves. Nature 179, 975–976.

Pearsall, D. M. (1979). The application of ethnobotanical techniquesto the problem of subsistence in the Ecuadorian Formative.Unpublished PhD thesis. Urbana: University of Illinois.

Pearsall, D. M. (1989). Paleoethnobotany: A Handbook of Proce-dures. London: Academic Press.

Pearsall, D. M., Piperno, D. R., Dinan, E. H., Umlauf, M., Zhao, Z.& Benfer, R.A. Jr (1995). Distinguishing rice (Oryza sativaPoaceae) from wild Oryza species through phytolith analysis:results of preliminary research. Economic Botany 49, 183–196.

Pearsall, D. M. & Trimble, M. K. (1984). Identifying past agricul-tural activity through soil phytolith analysis: a case study from theHawaiian Islands. Journal of Archaeological Science 11, 119–133.

Piperno, D. R. (1985). Phytolith analysis and tropical paleo-ecology:production and taxonomic significance of siliceous forms in newworld plant domesticates and wild species. Review of Palaeobotanyand Palynology 45, 185–228.

Piperno, D. R. (1988). Phytolith Analysis: An Archaeological andGeological Perspective. London: Academic Press.

Piperno, D., Bush, M. & Colinvaux, P. (1991). Paleoecologicalperspectives on human adaptation in central Panama, II: theHolocene. Geoarchaeology 6, 227–250.

Piperno, D. R. & Pearsall, D. M. (1993). Phytoliths in the reproduc-tive structures of Maize and Teosinte: implications for the study ofMaize evolution. Journal of Archaeological Science 20, 337–362.

Policard, A. (1923). La microincineration des cellules et des tissus.Protoplasma 7, 464–481.

Raeside, J. D. (1970). Some New Zealand plant opals. New ZealandJournal of Science 13, 122–132.

Richardson, W. D. (1920). The ash of dune plants. Science 1326,546–551.

Rosen, A. (1994). Identifying ancient irrigation: a new method usingopaline phytoliths from Emmer wheat. Journal of ArchaeologicalScience 21, 125–132.

Rovner, I. (1972). Note on a safer procedure for opal phytolithextraction. Quaternary Research 2, 591.

Rovner, I. (1983). Plant opal phytolith analysis: major advances inarchaeobotanical research. In (M. B. Schiffer, Ed.) Advances inArchaeological Method and Theory Vol. 6. London: AcademicPress.

Schelenberg, H. C. (1908). Wheat and barley from the NorthKurgan, Anau. In (R. Pumpelly, Ed.) Explorations in TurkestanVol. 3. Washington: Carnegie Institution, pp. 471–473.

Stewart, W. D. & Arthur, J. M. (1935). An improved standardizedmethod for ashing of plant material. American Journal of Botany22, 905.

886 J. F. Parr et al.

Twiss, P. C., Suess, E. & Smith, R. M. (1969). Morphologicalclassification of grass phytoliths. Soil Science Society of AmericaProceedings 33, 109–115.

Uber, F. M. (1940). Microincineration and ash analysis. The Botani-cal Review 6, 204–226.

Wang, S. S. & Hill, R. S. (1995). Phytolith analysis in leaves ofextant and fossil populations of Nothofagus subspecies Lophozo-nia. Australian Systematic Botany 8, 1055–1065.

Whang, S. S., Kim, K. & Hess, W. M. (1998). Variation of silicabodies in leaf epidermal long cells within and among seventeenspecies of Oryza (Poaceae). American Journal of Botany 85,461–466.

Wilson, S. M. (1985). Phytolith analysis at Kuk, an early agriculturalsite in Papua New Guinea. Archaeology in Oceania 20, 90–97.

Zimmerman, A. (1901). Botanical Microtechnique. Translated fromthe German by J. E. Humphrey. New York: Henry Holt.