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Experimental Attachment of Sediment Particles to Invertebrate Eggs and the Preservation of Soft-Bodied Fossils

Posted on: Saturday, 4 September 2004, 06:00 CDT

Clay minerals can be an important agent in the fossilization of soft tissues, notably in the Ordovician Soom Shale of South Africa and the Cambrian Burgess Shale of Canada. The replication of morphology has been attributed to adsorption of pre-existing clay minerals, or direct precipitation of authigenic clays onto tissues. Attachment of quartz and kaolinite to the surface of lobster eggs demonstrates experimentally for the first time that soft tissues could fossilize in pre-existing minerals. However, the eggs became coated only in the presence of metabolizing bacteria. This experimental approach could be used to explore why Burgess Shale- type preservation declined after the mid-Cambrian.

Keywords: Soom Shale, Burgess Shale, fossilization, mineralization, clay.

The majority of the fossil record consists of biomineralized or sclerotized remains as they have a high preservation potential. However, fossils of soft-bodied metazoans are relatively common and are an important source of palaeontological data. The preservation of soft tissues provides more detailed morphological information on extinct organisms and thus important information on evolutionary pathways, a more complete record of ancient community ecology and evidence of the factors controlling fossilisation. The fossilization of soft tissues normally requires the replication of their morphology by rapid in situ growth of minerals such as apatite, calcite and pyrite (Briggs 2003). Recent discoveries have demonstrated that replication in clay minerals, although rarer, is also important. The most remarkable example is the three- dimensionally preserved soft tissues of the giant conodont animal Promissum pulchrum, eurypterids and other arthropods from the Upper Ordovician Soom Shale of South Africa (Gabbott et al. 2001). In this case the surrounding sediment matrix is dominated by detrital quartz and illite along with various other clay minerals (e.g. kaolinite and chlorite) (Gabbott 1998). Clay minerals are also involved in the preservation of other biotas, most notably the Cambrian Burgess Shale, where different clay minerals occur on different soft tissues (Orr et al. 1998). Likewise they occur on crustaceans in the Carboniferous Coal Measures at Castlecomer in eastern Ireland (Orr & Briggs 1999) and insects in the Triassic deposits of Virginia (Fraser et al. 1996).

Gabbott (1998) proposed a model for the preservation of the Soom Shale fossils that involved attracting and adsorbing colloidal clay minerals onto the surface of organic substrates. However, on further consideration, Gabbott et al. (2001) favoured direct replacement of the soft tissues by authigenic illite (or a dioctahedral precursor), where the ions Si, A1 and K. were derived by dissolution of the surrounding sediment in a reduced pH environment. Orr et al. (1998) considered that either of these mechanisms might explain the preservation of the Burgess Shale fossils. Petrovich (2001), however, argued that preservation by the attachment of pre-existing minerals was unlikely because they would lack the surface configuration to 'dock' in the right orientation. Butterfield (1990, 1995) suggested a major role for clay minerals in preserving the Burgess Shale fossils by incapacitating autolytic and microbial digestive enzymes, which led to decay inhibition and the stabilization of structural polymers. In contrast, Petrovich (2001) argued that bacteria living in finegrained sediments were unlikely to employ extracellular enzymes that were affected in this way because they would be ineffective. However, investigations of marine sediments have shown that labile organic matter is stabilized by sorption onto mineral surfaces (Mayer 1993; Keil et al. 1994). Here it is demonstrated for the first time experimentally that the converse can also occur: mineral particles can become attached to organic matter, providing one possible mechanism to explain the preservation of softbodied fossils like those in the Burgess Shale and Soom Shale.

Method. The experiments (Table 1) were designed to determine some of the controls on the replication of soft tissue morphology in pre- existing mineral particles. For this preliminary investigation a simple substrate, lobster eggs, was selected. These eggs have been successfully used in experiments to investigate mineralization in calcium carbonate (Martin et al. 2003). The outer layer of a fresh lobster egg shows the remains of the funiculus (stalk) that once attached it to the female (Fig. 1a). Fresh eggs (100 eggs equivalent to 149mg dry mass) of the European lobster (Homarus gammarus L), obtained from the National Lobster Hatchery, Padstow, UK, were used in each experiment. They were shed naturally or accidentally rubbed off when moving gravid females during standard hatchery procedures. The eggs (including those to be used in non-sterile experiments) were placed in a 5% solution of the antiseptic chlorhexidine gluconate for 3 min to sterilize the surface, and then rinsed in sterile artificial seawater. They were placed in vials and covered by a slurry of thoroughly mixed sediment obtained from the top 100 mm in the Tamar Estuary, UK (Boschker el al. 1998). Weakly buffered, sterile (where applicable), oxic artificial seawater (4 m1) at pH 7.2 (Martin et al. 2003) was added to each vial to make the total volume up to c. 8 m1, before they were crimp sealed to prevent inward oxygen diffusion and incubated at 15 1 C for 31 days. Some experiments were incubated at 1 1 C to reduce bacterial metabolism as much as possible without freezing the medium. Two vials were used for each treatment and the experiment was run on two separate occasions. Sediment was sterilized, where required, by autoclaving at 121 C, 15p.s.i. for 30 min, allowing it to cool in a sealed container for 1 day, then re-autoclaving on the third day and cooling before use. Standard aseptic techniques were used where sterile incubation was required (i.e. all the equipment was sterilized and manipulations were performed in a laminar flow hood).

Table 1. Final pH of sediment surrounding the eggs and degree of particle attachment after incubation

Fig. 1. Scanning electron micrographs of lobster eggs, (a) Fresh egg. (b) After incubation in sterile sediment at 15 C for 31 days, showing no particles attached Io the surface, (c) After incubation in non-sterile sediment at 15 C for 31 days, showing particles attached to the surface, (d) Surface of egg in (c), showing no evidence of bacteria in the particle matrix, (e) After incubation in non-sterile sediment at 1 C for 31 days, showing no particles attached to the surface, (f) After incubation in non-sterile sediment at 15 C for 10 months, showing a complete covering of particles attached to the surface, (g) Surface of egg in (f), showing no evidence of bacteria in the particle matrix, (h) Sliced egg after incubation in non-sterile sediment at 15 C for 10 months, showing the layer of particles that surrounds the egg (1) and the unmineralized internal contents of the egg (2). Scale bar represents 500 m in (a)-(c), (c) and (f); 30 m in (d); 50 m in (g); 100 m in (h).

After incubation the pH at the bottom of the vial, amongst the eggs, was measured with a microelectrode (Diamond General, Ann Arbor, MI). The eggs were gently washed under a flow of artificial seawater until the rinse was clear of sediment particles. The eggs were prepared for scanning electron microscopy using the method of Sagemann et al. (1999). At least six eggs from each of the five treatments (Table 1) were carbon coated before scanning on a Hitachi S-3500N instrument with an accelerating voltage of 12-15kV The elemental composition of the particles surrounding the eggs was determined using an energy-dispersive X-ray (EDX) detector and XRD was performed on the particles attached to the eggs using a Philips PW1710 diffractometer with CuK [alpha]-radiation at 35 kV and 40 mA to determine the mineral composition.

Results. Where bacterial metabolism was not inhibited, cither by sterilizing the sediment or lowering the temperature to 1 C, the pH in the vials fell to 6.9 and the sediment went black, reflecting the activity of sulphate-reducing bacteria via iron sulphide formation (Table 1, treatments D and E). Otherwise the pH remained at 7.2 or fell slightly to 7.1 and the sediment remained grey, indicating the absence of bacterial sulphate reduction (Table 1, treatments A-C).

Where the eggs were incubated in sterile sediment at 15 C (Table 1, treatment B) for 31 days no particles remained attached to the outer envelope after washing (Fig. 1b); the same applied where eggs were incubated in sterile sediment at 1 C (Table 1, treatment A). Where incubation was performed with non-sterile sediment at 15 C (Table 1, treatment D) particles of sediment from 1 to 20 m in size covered the outer egg envelope after washing (Fig. 1c and d). Sustained incubation for 10 months (Table 1, treatment E) resulted in a greater density of particles to a thickness of c. 15 m attached to the surface (Fig. 1f and g), but no minerals were observed inside the egg, which remained unaffected (Fig. 1h\). In contrast, where incubation was performed with non-sterile sediment at just 1 C, with consequent reduction in bacterial activity (Table 1, treatment C), no particles attached to the outer envelope (Fig. 1e). EDX analysis of the sediment particles attached to the eggs revealed large aluminium and silicon peaks, suggesting the presence of clay minerals. Semi-quantitative analysis of the XRD data showed that the particles surrounding the eggs were composed of quartz and kaolinite. Neither the composition, as revealed by XRD, nor the size and morphology of the particles differed from those of the sediment that was added to the vials.

Discussion. The particles attached to the egg are the same composition, size and morphology as those in the sediment. This demonstrates that attachment of pre-existing particles in the presence of active bacteria is the likely mechanism of mineral coat formation, rather than authigenic precipitation. In these experiments the decrease in pH was similar to that generated by decay at 15C (Table 1, treatments D and E) in other decay experiments (Briggs & Kear 1994; Hof & Briggs 1997). This probably reflects the release of H^sub 2^S, CO2 and fatty acids produced during decomposition (Briggs & Kear 1994). Where the sediment was sterile, or bacterial metabolism was slowed by a lowered temperature, the pH remained virtually unchanged, reflecting the lack of decomposition. The reduced pH may have facilitated direct sorption of the sediment particles, in particular clays, onto the surface of the eggs (see Gabbott 1998), although any effect would probably be minimal considering the small pH decrease. Sediment particle attachment to the egg envelope, therefore, was probably a direct result of the presence of active bacteria. Where bacteria were absent, or their activity was inhibited by a lowered temperature, sediment particles did not attach to the eggs.

These experiments show that quartz and clay particles can attach to the surface of the organic material that makes up the egg envelope (contra Petrovich 2001). This occurred within 31 days and the outer envelope became coated in sediment particles to a thickness of c. 15 m within 10 months (Fig. 1f). It is not known whether the attachment of these particles inhibited further organic decomposition; to determine this would require longer-term experiments. The egg envelope coated by the sediment particles is essentially uniform in biochemical composition. Variation in composition, in the soft tissues of a carcass, for example, might influence the attachment of different minerals or release of certain ions, resulting in contrasts in the composition of different parts of the resultant fossil, and between the fossil and the matrix (Orr et al. 1998). Small clay particles (1 nm1 m ), such as those that preserve the Soom Shale fossils, have the potential to preserve soft tissue morphology with a fidelity as high as that characteristic of apatite (Gabbott 1998; Briggs 2003). The particles involved in our experiments (1-20 m), in contrast, would yield less information at this scale.

Presumably, the presence of actively metabolizing bacteria made the surface of the eggs more amenable to particle attachment. None the less, bacterial cells were never observed on or within the sediment particles (Fig. 1d and g). However, bacteria are rarely preserved in laboratory decay experiments except where they themselves become mineralized (Sagemann et al. 1999). Thus we cannot eliminate the possibility that bacteria, or extracellular polysaccharides produced by them, acted as an attachment site for the sediment particles. Also, we cannot rule out the possibility that direct precipitation of clays, in particular, may be important in certain circumstances (e.g. Konhauser et al. 1993; Gabbott et al. 2001; Kim et al. 2004). Obviously, coating by sediment particles represents only the first stage in the fossilization of soft tissues. Both the envelope and content of the egg might subsequently decay. Alternatively, stabilization of the outside of the egg by the attachment of sediment particles might prevent collapse and facilitate further authigenic mineralization of the interior (Xiao & Knoll 2000; Martin et al. 2003). Survival of the potential fossil requires cementation of the surrounding sediment matrix and the covering of particles may be altered subsequently through diagenesis.

The initial stage of preservation of soft tissues in sediment particles (i.e. attachment to an organic substrate) will depend on a number of variables including pH, mineral composition, available exchange cations and tissue composition (see Buttcrfield 1995). Further experimental investigation of these variables is necessary to determine how readily different minerals attach to organic matter in different circumstances. This would provide an experimental test, for example, of the hypothesis that long-term changes in clay mineralogy were important in determining the distribution of Burgess Shale-type preservation through time (Butterfield 1995, 2003) and in largely closing that taphonomic window (sensu Allison & Briggs 1991) after the Mid-Cambrian.

This work was funded by Natural Environment Research Council grant GR3/12465 to D. E. G. B. and R. J. P. while all the authors were at the University of Bristol. We thank R Midglcy from the National Lobster Hatchery for providing the lobster eggs, and J. Charmant (University of Bristol) and A. Oldroyd (University of Cardiff) for XRD analysis. S. E. Gabbott and D. Pirrie provided helpful comments.

References

ALLISON, P.A. & BRIGGS, D.E.G. 1991. Taphonomy of non- mineralized tissues. In: ALLISON, P.A. & BRIGGS, D.E.G. (eds) Taphonomy: Releasing the Data Locked in lhe Fossil Record. Plenum, New York, 25-70.

BOSCHKER, H.T.S., NOLD, S.C., WELLSUUUY, P., DE GRAAF, W., PEL, R., PARKES. R.J. & CAPPENBERG, T.E. 1998. Direct linking of microbial populations to specific biogeochemical processes by 13C- labelling of biomarkers. Nature, 392, 801-805.

BRIGGS, D.H.G. 2003. The role of decay and mineralization in the preservation of soft-bodied fossils. Annual Review of Earth and Planetary Sciences, 31, 275 301.

BRIGGS, D.E.G. & KEAR, A.J. 1994. Decay and mineralization of shrimps. Palaios, 9, 431-456.

BUTTERFIELD, N.J. 1990. Organic preservation of non-mineralizing organisms and the taphonomy of the Burgess Shale. Paleobiology, 16, 272-286.

BUTTERFIELD, N.J. 1995. Secular distribution of Burgess-Shale- type preservation. Lethaia, 28, 1-13.

BUTTERFIELD, N.J. 2003. Exceptional fossil preservation and the Cambrian Explosion. Integrative and Comparative Biology, 43, 166- 177.

FRASER, N.C., GRIMALDI, D.A., OLSEN, P.E. & AXSMITH, B. 1996. A Triassic Lagersttte from eastern North America. Nature, 380, 615- 619.

GABBOTT, S.E. 1998. Taphonomy of the Ordovician Soom Shale Lagersttte: an example of soft tissne preservation in clay minerals. Palaeontology, 41, 631-667.

GABBOTT, S.E., NORRY, M.J., ALDRIDUE, R.J. & THERON, J.N. 2001. Preservation of fossils in clay minerals; a unique example from the Upper Ordovician Soom Shale, South Africa. Proceedings of the Yorkshire Geological Society, 53, 237-244.

HOF, C.H.J. & BRIGGS, D.E.G. 1997. Decay and mineralization of mantis shrimps (Stomatopoda: Crustacea)-a key to their fossil record. Palaios, 12, 420-438.

KEIL, R.G., MONTLUCON, D.B., PUAHL, E.G. & HEDGES, J.I. 1994. Sorptive preservation of labile organic matter in marine sediments. Nature, 370, 549-552.

KIM, J., DONG, H., SHABAUGH, J., NHWHU., S.W. & CBERL, D.D. 2004. Role of microbes in the smectite-to-illite reaction. Science, 303, 830 832.

KONHAUSER, K.O., FYFE, W.S., FERRIS, F.G. & BEVBRIDGE, T.J. 1993. Metal sorption and mineral precipitation by bacteria in two Amazonian river systems: Rio Solimoes and Rio Negro, Brazil. Geology, 21, 1 103-1106.

MARTIN, D., BRIGGS, D.E.G. & PARKHS, R.J. 2003. Experimental mineralization of invertebrate eggs and the preservation of Neoprotcrozoic embryos. Geology, 31,39-42.

MAYER, I.M. 1993. Surface-area control of organic carbon accumulation in continental shelf sediments. Geochimica et Costnochiniica Ada, 58, 1271-1284.

ORR, PJ. & BRIGGS, D.E.G. 1999. Exceptionally preserved conchostracans and other crustaceans from the upper Carboniferous of Ireland. Special Paper* in Palaeontology, 62, 1-68.

ORR, P.J., BRIGGS, D.E.G. & KEARNS, S.L. 1998. Cambrian Burgess Shale animals replicated in clay minerals. Science, 281, 1173-1175.

PETROVICH, R. 2001. Mechanisms of fossilization of the soft- bodied and lightly armored faunas of the Burgess Shale and of some other classical localities. American Journal of Science, 301, 683- 726.

SAGFMANN, J., BALE, S.J., BRIGGS, D.E.G. & PARKES, R.J. 1999. Controls on the formation of authigenic minerals in association with decaying organic matter: an experimental approach. Geochimica et Cosmochimica Ada, 63, 1083-1095.

XIAO, S. & KNOU., A.H. 2000. Phosphatized animal embryos from the Neoproterozoic Doushanluo Formation at Wcng'an, Guizhou, South China. Journal of Paleontology 74, 767-788.

Received 27 October 2003; revised typescript accepted 14 April 2004.

Scientific editing by Duncan Pirrie

DEREK MARTIN1, DEREK E. G. BRIGGS2 & R. JOHN PARKES1

1 School of Earth, Ocean and Planetary Sciences, Cardiff University, Main Building, PO Box 914, Park Place, Cardiff CF10 3YE, UK (e-mail: dmartin@earth.cf.ac.uk)

2 Department of Geology and Geophysics, Yale University, PO Box 208109, New Haven, CT 06520-8109, USA

Copyright Geological Society Publishing House Sep 2004

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