Plant macrofossil analysis of Holocene alluvium, with special reference to the
Lower Thames Basin
Andrew Stephen Fairbairn
2001
Submitted as part of the fulfilment for
the degree of Doctor of Philosophy
Institute of Archaeology
I Abstract
Alluvium is an important archaeological and palaeoenvironmental resource in lowland Britain. The research presented here develops plant macrofossil analysis of alluvial facies, with special emphasis on the depositional and natural environments of the Lower Thames Basin. Plant
macrofossil analysis is a poorly developed area of alluvial research, usually limited to superficial description of the fossils seen in section, or detailed analysis of a narrow suite of remains. A comprehensive, quantitative method of macrofossil analysis using counts and cover abundance scores is developed. Identification criteria for several groups of macrofossils are presented, including leaves, rootlets and epidermis.
Potential macrofossil incorporation was investigated at eight wetland and alluvial sites, including saltmarsh, wet woodland and herb fen environments. Macrofossil collections were compared to extant vegetation and subject to multivariate analysis. The results showed that
macrofossil assemblages produce spatially and temporally precise data of plant presences, although spatial and temporal fidelity varies in different depositional environments and between plant taxa. Vegetation dominants were favoured in the assemblages of all classes of
macrofossils, with bulky Monocotyledons and Therophytes favourably preserved and sparsely distributed taxa, such as rosette plants, less well favoured. The depositional environment and position in relation to environmental gradients were also found to affect macrofossil
composition. Multiple approaches to macrofossil analysis using a wide range of macrofossils were found to produce improved interpretations. The value of different macrofossil classes and
occurrences of the major observed taxa in alluvial sediments are discussed.
The method was applied to samples from the Medway River at Chatham. Vegetation history, hydrology and traces of human disturbance are discussed from 7000BP to 2000BP.
Analysis showed a gradual increase in human disturbance over time, development of a distinctive human-influenced upper salt marsh flora from 3000BP and supports the trend across
3.
Acknowledgements
I thank Dr. Jon Hather and Tony Barham for supervising this project and the GSF staff (Chris Pine, Vaughan Williamson, Dr. Martin Bates) who not only recovered the Medway Tunnel material, but provided some of the stimulus for the project. Thanks also to to the wardens of the nature reserves, owned by the RSPB, English Nature,
Suffolk Wildlife Trust, Kent Wildlife Trust and Norfolk Naturalists Trust, for their patience and willing help. David Eagle gave permission for access to Stonemarsh. Phil Austin, Dr. Louise Martin, Helen Wickstead, Sophie Seal and many other members of the Institute of Archaeology provided social support and discussion. Sandra Bond and Harry Pratt provided invaluable technical support, without which the identification work would have been impossible. Dr. Paul Hughes and Prof. Keith Barber provided advice
4
Table of Contents
Page
Abstract
2
Acknowledgements
3
Table of Contents
4
List of figures
12
List of tables
18
List of plates 20
1 Introduction 28
2 Plant macrofossil analysis in archaeological and palaeoenvironmental 33 investigations of Holocene alluvium, with special reference to the Lower
Thames Basin
2.1 Nature and distribution of Holocene alluvium
33
2.2 Rivers and estuaries as sedimentary basins 34
2.2.1 Hydrological factors
34
2.2.2 Sedimentation 35
2.2.3 Morphology 36
2.2.4 Riverine and estuarine plant ecology
38
2.2.4.1 Environmental influences 38
2.2.4.2 Rivers 38
2.2.4.3 Estuaries 39
2.2.5 Depositional environments, erosional environments and 40
facies
2.2.5.1 Rivers
40
2.2.5.2 Estuaries 41
2.3 Plant macrofossil analysis
42
2.3.1 Plant macrofossils in geology and archaeology 42
2.3.2 Methods and techniques 43
2.3.2.1 Plant macrofossils in facies descriptions
43
2.3.2.2 Sampling and recovery
44
2.3.2.3 Identification 45
2.3.2.4 Recording and presentation of macrofossil 48 assemblages
5
2.3.3 An interpretative framework 50
2.3.4 Taphonomy 51
2.3.4.1 Biological considerations 52
2.3.4.2 Necrology 53
2.3.4.3 Biostratinomy 58
2.3.4.4 Diagenesis 60
2.3.4.5 Actualistic research 60
2.3.4.6 Temporal and spatial-averaging 62
2.3.4.7 Implications
63
2.4 Palaeoenvironmental and palaeoecological applications 63 2.4.1 Interpreting alluvium in space and time 63
2.4.2 Channel and floodplain development
65
2.4.3 Palaeohydrology 65
2.4.4 Palaeoecology
and vegetation history
66
2.4.5 Local terrestrial environmental change 68
2.4.6 Relative sea-level change. 69
2.4.7 Climatic change.
70
2.5 The archaeology of alluvium
71
2.5.1 Alluvium as an archaeological resource 71 2.5.2Preserved sites, features and structures 72 2.5.3 Rivers, floodplains, estuaries and saltmarshes in the 74 human past
2.5.4 Human-wetland interactions in the Lower Thames
75
2.6 Plant macrofossil analysis of alluvial facies: Potential and 77 problems
3 Laboratory methods
79
3.1 Classification 79
3.1.1 Moss and liverwort (Bryophyta) macrofossils 82
3.1.2 Aerial structures 82
3.1.2.1 Woody stems
82
3.1.2.2 Non-woody aerial stems
83
3.1.2.3 Buds and bud-scales 83
3.1.2.4 Leaves 84
6
3.1.2.5.1 Stipules
85
3.1.2.5.2 Thorns 85
3.1.2.5.3 Prickles 85
3.1.2.5.4 Scales 85
3.1.3 Underground structures 85
3.1.3.1 Non-woody underground stems 85
3.1.3.2 Rootlets
86
3.1.4 Dispersal and reproductive structures 86
3.1.4.1 Spores and sporangia
86
3.1.4.2 Seeds and fruits 87
3.1.4.2.1 Seeds and similar structures 87 3.1.4.2.2 Bracts and similar structures 87
3.1.5 General 88
3.1.5.1 Tissues 88
3.1.5.1.1 Epidermis and cuticles
88
3.1.5.1.2 Vascular strands 88
3.1.5.1.3 Wood 88
3.1.5.1.4 Unidentified vegetativetherbaceous
88
matter
3.1.5.2 Indeterminate
88
3.2 Identification 89
3,2.1 Selected Taxa 89
3,2.2 Reference specimen preparation
92
3.2.2.1 Specimen collection and pre-treatment 92
3.2.2.2 Clearing. 93
3.2.2.3 Staining and mounting methods
94
3.2.2.4 Recording methods
95
3.2.3 Identification criteria 98
3.2.3.1 Rootlets
98
3.2.3.2 Sphenophyta
leaf-sheaths
and epidermis
100
3.23.3 Filicophyta leaves and epidermis
102
3.2.3.4 Dicotyledonae leaves
105
3.2.3.5 Monocotyledonae leaves and epidermis 112
I
3.3 Quantification and recording 115
3.3.1 The cover abundance method 116
3.3.2 Tests 117
3.3.3 Comparability of cover abundance
values
117
3.3.4 Evaluation 120
4. An investigation of plant macrofossil taphonomy in modern alluvial
122
environments
4.1 Site selection 122
4.2 Field and laboratory methods
123
4.2.1 Sample point selection 123
4.2.2 Sediment sampling 125
4.2.3 Sediment description 126
4.2.4 Vegetation recording. 126
4.2.5 Laboratory preparation, description and analysis of
127
macrofossils
4.2.6 Sediment tests 129
4.3 Stonemarsh 131
4.3.1 Location and topography 131
4.3.2 Vegetation and surface litter 131
4.3.3 Sampling
131
4.3.4 Sediments 135
4.3.5 Incorporation, sources and preservation of macrofossils 135
4.3.6 Sample size effects
139
4.3.7 Quantitative analysis 139
4.3.8 Differences between depositional environments 145
4.3.9 Vegetation representation
145
4.3.10 Sub-surface samples from sediment blocks 147
4.4 Borstal Marsh 147
4.4.1 Location and topography 147
4.4.2 Vegetation and surface litter
147
4.4.3 Sampling
151
4.4.4 Sediments
151
4.4.5 Incorporation, sources and preservation of macrofostils 152
8'
4.4.7 Vegetation representation 159
4.5. Angel Marsh, Walberswick 160
4.5.1 Location and topography 160
4.5.2 Vegetation and surface litter
160
4.5.3 Sampling 164
4.5.4 Sediments 164
4.5.5 Incorporation, sources and preservation of macrofossils
165
4.5.6 Sample size effects
166
4.5.7 Quantitative analysis 176
4.5.8 Vegetation representation 177
4.5.9 Sub-surface samples from sediment blocks 183
4.6 Snape Saltings 183
4.6.1 Location and topography 183
4.6.2 Vegetation and surface litter 185
4.6.3 Sampling
186
4.6.4 Sediments 186
4.6.5 Incorporation, sources and preservation of macrofossils 191
4.6.6 Sample size effects
204
4.6.7 Quantitative analysis 207
4.6.8 Differences in depositional environment 213
4.6.9 Vegetation representation 214
4.6.10 Sub-surface samples from sediment blocks 218
4.7. Burham Marsh 218
4.7.1 Location and topography 218
4.7.2 Vegetation and surface litter 218
4.7.3 Sampling
219
4.7.4 Sediments
219
4.7.5 Incorporation, sources and preservation of macrofossils 224
4.7.6 Sample size effects
235
4.7.7 Quantitative analysis 235
4.7.8 Differences in depositional environment
237
4.7.9 Vegetation representation
242
4.7.10 Sub-surface samples from sediment blocks 243
9
4.8.1 Location and topography
244
4.8.2 Vegetation and surface litter 244
4.8.3 Sampling 245
4.8.4 Sediments 245
4.8.5 Incorporation, sources and preservation of macrofossils 253
4.8.6 Sample size effects 255
4.8.7 Quantitative analysis
256
4.8.8 Vegetation representation 257
4.9. Wet woodlands at Bure Marshes and Wicken Fen
263
4.9.1 Location and topography 263
4.9.2 Vegetation and surface litter 266
4.9.3 Sampling 268
4.9.4 Sediments 268
4.9.5 Incorporation, sources and preservation of macrofossils 269
4.9.6 Sample size effects
277
4.9.7 Quantitative analysis 277
4.9.8 Differences in depositional environment 284
4.9.9 Vegetation representation
284
4.9.10 Sub-surface
samples from sediment Blocks
290
4.10. Synthesis
291
4.10.1 Macrofossil characterisation of different depositional 291 environments
4.10.1.1 Saltmarsh
environments
291
4.10.1.2 Freshwater environments 296
4.10.1.3 Transitions 297
4.10.1.4 Interpreting facies changes in time and space
298
4.10.2 Vegetation reconstruction 301
4.10.2.1 Spatial fidelity of macrofossil assemblages
301
4.10.2.2 Temporal fidelity of macrofossil assemblages
303
4.10.2.3 Taxonomic fidelity of macrofossil assemblages 304
4.10.2.4 Taxon presence 306
4.10.2.5 The quantitative reconstruction of vegetation
307
10
4.10.2.6 Alluvial macrofossil assemblages as a basis for 311
vegetation reconstruction
4.10.3 The usefulness of different types of macrofossils 314
4.10.4 Autotaphonomy of species 319
4.10.4.1 Tree species
319
4.10.4.2 Shrubs and sub-shrubs 324
4.10.4.3 Bulky Monocotyledon 325
4.10.4.4 Perennial Dicotyledon herbs and 328 Monocotyledon rosette plants
4.10.4.5 Therophytes 331
4.10.4.6 Climbers 333
5. Plant macrofossil analysis of alluvial facies from the River Medway,
334
Chatham, Kent
5.1 The site 334
5.2 Stratigraphy and archaeological evidence 335
5.3 Materials and Methods
339
5.3.1 Sample framework 339
5.3.1.1 Phase
I Section 16
339
5.3.1.2 Phase I Section 9 340
5.3.1.3 Phase I Section 11 340
5.3.1.4 Phase
I Section 6
340
5.3.1.5 Phase III Trench 7 341
5.3.1.6 Phase III Trench 6 341
5.3.2 Sampling 341
5.3.3 Sample preparation 342
5.3.4 Recording and identification 342
5.4 Results
342
5.4.1 Range of preserved macrofossils 342
5.4.2 Section 16 344
5.4.2.1 Macrofossil preservation
344
5.4.2.2 Analysis
358
5.4.2.3 Interpretation
362
5.4.3 Section 9 368
11
5.4.3.2 Analysis 368
5.4.3.3 Interpretation 372
5.4.4 Section 11 374
5.4.4.1 Macrofossil preservation 374
5.4.4.2 Analysis 378
5.4.4.3 Interpretation
379
5.4.5 Section 6 381
5.4.5.1 Macrofossil preservation 381
5.4.5.2 Analysis 382
5.4.5.3 Interpretation 388
5.4.6 Trench 7 391
5.4.6.1 Macrofossil preservation 391
5.4.6.2 Analysis 394
5.4.6.3 Interpretation 395
5.4.7 Trench 6
398
5.5 Major research themes 399
5.5.1 Vegetation cover 399
5.5.1.1 River and riverbank vegetation 399
5.5.1.2 Terrestrial vegetation 401
5.5.1.3 Vegetation change and stability
402
5.5.2 Changes in depositional environment and hydrology 403
5.5.3 Human activity and disturbance 405
5.5.4 Usefulness
of Method
407
6 Discussion 409
7 Conclusions 421
Bibliography
423
Appendix I- Root descriptions 456
Appendix 2- Dicotyledonae Leaf Characters 462
12
List of figures
Figure 2.1 Framework of plant macrofossil analysis 50
Figure 2.2 The relationship of palaeoecology and the component disciplines 52
of taphonomy based upon the time interval of the organisms under study
Figure 2.3 Raunkaier's life-form classification 55
Figure 2.4 Freshwater (glycophytic) and saltwater (halophytic) indicator 68
species,
including those of strong (euhalobic), medium (mesohalobic) and
weak (oligohalobic) salinity
Figure 3.1 Phylogenetic relationships in the plant kingdom
80
Figure 3.2 Staining and mounting procedure 95
Figure 3.3 Sample 04/116 Cover Abundance Trial 8 119
Figure 4.1 Figure 4.1 Sites sampled and described in Chapters 4 and 5 124 Figure 4.2 Map of Stonemarsh showing sample points 132 Figure 4.3 Stonemarsh organic content and % water by weight 135 Figure 4.4 Stonemarsh seed and species abundance data 137 Figure 4.5 Stonemarsh seed and species concentration data 137 Figure 4.6 Stonemarsh: Percentage of seeds from plants at set distances from 138
sample points
Figure 4.7 Stonemarsh
Area 1 Sample ubiquity data
138
Figure 4.8a Stonemarsh cumulative seed concentration for Block 3 141 Figure 4.8b Stonemarsh: Cumulative percentage plots for major non-seed 141
elements
Figure 4.9a Correspondence
analysis of seed data from Stonemarsh
142
Figure 4.9b Canonical correspondence analysis of seed data from Stonemarsh 142 Figure 4.1 Oa Correspondence analysis of non-seed data from Stonemarsh 143
Figure 4.1Ob Canonical correspondence
analysis of non-seed
data from
143
Stonemarsh
Figure 4.11 a Correspondence analysis of seed and non-seed data from 144
Stonemarsh
Figure 4.1 lb Canonical correspondence
analysis of seed and non-seed data
144
from Stonemarsh
Figure 4.12a Sites on the Medway river 148
Figure 4.12b Borstal Marsh sample points 148
13
Figure 4.14 Borstal Marsh seed and species concentration data 153 Figure 4.15 Borstal Marsh percentage of seeds at set distances of the sample 153
points
Figure 4.16 Borstal Marsh sample ubiquity data 154
Figure 4.17a Correspondence analysis of seed data from Borstal Marsh 156 Figure 4.17b Canonical correspondence analysis of seed data from Borstal 156
Marsh
Figure 4.18a Correspondence
analysis of non-seed data from Borstal Marsh
157
Figure 4.18b Canonical correspondence
analysis of non-seed
data from
157
Borstal Marsh
Figure 4.19a Correspondence analysis of seed and non-seed data from Borstal 158
Marsh
Figure 4.19b Canonical correspondence analysis of seed and non-seed data 158 from Borstal Marsh
Figure 4.20 Angel Marsh location map and sampling points 161 Figure 4.21 Angel Marsh organic content and % water 164 Figure 4.22 Angel Marsh seed and species concentration data 171 Figure 4.23 Angel Marsh percentage of seeds from plants at set distances 172
from the sample points
Figure 4.24a Angel Marsh sample ubiquity data Transect 1
173
Figure 4.24b Angel Marsh sample ubiquity data Transect 2 173 Figure 4.25a Angel Marsh Block 1: Cumulative seed data for main sample 174
components
Figure 4.25b Angel Marsh Block 2: Cumulative seed data for main sample 174 components
Figure 4.26a Angel Marsh Block 1: Cumulative non-seed data for main
175
sample components
Figure 4.26b Angel Marsh Block 2: Cumulative non-seed data for main 175
sample components
Figure 4.27a Angel Marsh correspondence
analysis of combined seed and
178
non-seed
data from block samples
Figure 4.27b Angel Marsh canonical correspondence analysis of combined 178 seed and non-seed data from block samples
14
Figure 4.28b Canonical correspondence analysis of seed data from Angel 179 Marsh
Figure 4.29a Correspondence analysis of non-seed data from Angel Marsh 180 Figure 4.29b Canonical correspondence analysis of non-seed data from Angel 180 Marsh
Figure 4.30a Correspondence
analysis of seed and non-seed
data from Angel
181
Marsh
Figure 4.30b Canonical correspondence analysis of seed and non-seed data 181
from Angel Marsh
Figure 4.31 Snape Saltings location map and sample point information 184 Figure 4.32 Snape Saltings organic content and % water 190
Figure 4.33 Seed and species concentration data for transect samples
198
Figure 4.34a Snape Saltings seed and species abundance data for Blocks 199
Figure 4.34b Snape Saltings seed and species concentration data for Blocks
199
Figure 4.35 a Snape Saltings percentage of seeds from set distances from the
200
sample point
Figure 4.35 b Snape Saltings percentage of seeds from set distances from the 201
sample point
Figure 4.36 a Snape Saltings sample ubiquity data 202
Figure 4.36b Snape Saltings sample ubiquity data
203
Figure 4,37a Snape Saltings Block 1 cumulative seed data for major taxa 205 Figure 4.37b Snape Saltings Block3 cumulative seed data for major taxa 205
Figure 4.38a Snape Saltings Block 1 cumulative non-seed data for major taxa
206
Figure 4.38b Snape Saltings Block 3 cumulative non-seed data for major taxa 206 Figure 4.39a Snape Saltings correspondence analysis of combined seed and 209
non-seed data from block samples
Figure 4.39b Snape Saltings canonical correspondence analysis of combined 209
seed and non-seed
data from block samples
Figure 4.40a Correspondence analysis of seed data from Snape Saltings 210
Figure 4.40b Canonical correspondence
analysis of seed data from Snape
210
Saltings
Figure 4.41 a Correspondence analysis of non-seed data from Snape Saltings 211 Figure 4.41b Canonical correspondence analysis of non-seed data from Snape 211
15
Figure 4.42a Correspondence
analysis of seed and non-seed
data from Snape
212
Saltings
Figure 4.42b Canonical correspondence
analysis of seed and non-seed
data
212
from Snape Saltings
Figure 4.43 Burham Marsh location map and sample points 220 Figure 4.44 Burham Marsh sample organic content and % water 223
Figure 4.45 Burham Marsh Transect sample seed and species concentration
229
Figure 4.46a Burham Marsh Block sample seed and species abundance data 230
Figure 4.46b Burham Marsh Block sample seed and species concentration
230
data
Figure 4.47 Burham Marsh: Percentage of seeds from set distances from the 231 sample point
Figure 4.48a Burham Marsh ubiquity data 232
Figure 4.48b Burham Marsh ubiquity data 232
Figure 4.49a Burham Marsh Block 1 cumulative seed data for main taxa
233
Figure 4.49b Burham Marsh Block 3 cumulative seed data for main taxa 233 Figure 4.50a Burham Marsh Block 1 cumulative non-seed data for main taxa 234 Figure 4.50b Burham Marsh Block 3 cumulative non-seed data for main taxa 234 Figure 4.51a Burham Marsh correspondence analysis of seed and non-seed 238
data from block samples
Figure 4.5 lb Burham Marsh canonical correspondence analysis of seed and 238 non-seed data from block samples
Figure 4.52a Burham Marsh correspondence
analysis of seed data from
239
Transect samples
Figure 4.52b Burham Marsh canonical correspondence analysis of seed data 239
from Transect samples
Figure 4.53a Burham Marsh correspondence
analysis of non-seed
data from
240
Transect samples
Figure 4.53b Burham Marsh canonical correspondence analysis of non-seed 240
data from Transect samples
Figure 4.54a Burham Marsh correspondence
analysis of combined seed and
241
non-seed data from Transect samples
Figure 4.54b Durham Marsh canonical correspondence analysis of combined 241
16
Figure 4.55 Hickling Broad location map and sample points 246 Figure 4.56 Hickling Broad organic and water content 245 Figure 4.57 Hickling Broad Block 1 cumulative seed data for main taxa 250 Figure 4.58 Hickling Broad Block 1 cumulative non-seed data for main taxa 251 Figure 4.59a Hickling Broad seed and species abundance 252 Figure 4.59b Hickling Broad seed and species concentration 252
Figure 4.60 Hickling Broad: Percentage
of seeds
from set distances from the
254
sample point
Figure 4.61 Hickling Broad sample ubiquity data
255
Figure 4.62a Hickling Broad correspondence analysis of seed data 258 Figure 4.62b Hickling Broad canonical correspondence analysis of seed data 258 Figure 4.63a Hickling Broad correspondence analysis of non-seed data 259 Figure 4.63b Hickling Broad canonical correspondence analysis of non-seed 259
data
Figure 4.64a Hickling Broad correspondence
analysis of seed and non-seed
260
data
Figure 4.64b Hickling Broad canonical correspondence analysis of seed and 260
non-seed
data
Figure 4.65 Bure Marsh location map and sample points 264
Figure 4.66 Wicken Fen location and sample points
265
Figure 4.67 Bure marsh and Wicken Fen organic and water data 269 Figure 4.68a Wicken Fen and Bure Marsh seed, seed and bract and species 272
abundance
Figure 4.68b Wicken Fen and Bure Marsh seed and species concentration 272 Figure 4.69 Wicken Fen and Bure Marsh: Percentage of seeds from set 273
distances from the sample point
Figure 4.70a Wicken Fen and Bure Marsh: % of arboreal and non-arboreal 273 taxa (minus Filicales) in seed assemblages
Figure 4.70b Wicken Fen and Bure Marsh: % of seeds from arboreal and non-
274
arboreal species
(all seeds)
Figure 4.71 Wicken Fen and Bure Marsh sample ubiquity data 274
Figure 4.72a Bure Marsh Block 1 cumulative seed data for main taxa
279
Figure 4.72b Bure Marsh Block 1 cumulative non-seed data for main taxa 279
17
Figure 4.72c Bure Marsh Block 1. cumulative seed data for canopy taxa 280 Figure 4.72d Bure Marsh Block 1 cumulative seed data for groundstorey taxa 280 Figure 4.73a Wet Woodland correspondence analysis of seed data 281 Figure 4.73b Wet Woodland canonical correspondence analysis of seed data 281
Figure 4.74a Wet Woodland correspondence
analysis of non-seed
data
282
Figure 4.74b Wet Woodland canonical correspondence
analysis of non-seed
282
data
Figure 4.75a Wet Woodland correspondence analysis of seed and non-seed 283
data
Figure 4.75b Wet Woodland canonical correspondence
analysis of seed and
283
non-seed
data
Figure 4.76a Comparative DOMIN scores of standing vegetation cover 286
abundance
(CA) and seed data for the main arboreal taxa
Figure 4.76b Comparative DOMIN scores of standing vegetation cover
287
abundance
(CA) and bud-scale data for the main arboreal taxa
Figure 4.76c Comparative DOMIN scores of standing vegetation cover
288
abundance
(CA) and bract data for the Betulaceae taxa
Figure 4.77a Comparative ubiquity data of seeds from arboreal and 321
Monocotyledon taxa present in modern surface samples
Figure 4.77b Comparative ubiquity data of seeds from herbaceous
322
Dicotyledon taxa and Monocotyledon rosette taxa present in modern surface
samples
Figure 5.1 Plan and section of the Medway Tunnel site eastern approach
336
Figure 5.2 Schematic
profile of the sediments
at the Medway Tunnel site with
337
relevant sections numbered (after Bates and Bates 2000)
Figure 5.3 Medway Tunnel Section 16 LOI figures 355
Figure 5.4 Medway Tunnel Section 16 Seed and species concentration
356
Figure 5.5 Medway Tunnel Section 16 CA diagram for all macrofossils 359 Figure 5.6 Medway Tunnel Section 16 CA of non-seed macrofossil data 360
Figure 5.7 Medway Tunnel Section 9 Seed and species concentration data
370
Figure 5.8 Medway Tunnel Section 9 CA diagram of all macrofossil data
370
Figure 5.9 Medway Tunnel Section 9 LOI data 371
Figure 5.10 Medway Tunnel Section 11 LOI data 377
18
Figure 5.12 Medway Tunnel Section 11 CA diagram of all macrofossil data 378
Figure 5.13 Medway Tunnel Section 6 LOI data 386
Figure 5.14 Medway Tunnel Section 6 seed and species concentration data 386 Figure 5.15 Medway Tunnel Section 6 CA of all macrofossil data 387 Figure 5.16 Medway Tunnel Trench 7 seed and species concentration data 393
Figure 5.17 Medway Tunnel Trench 7 LOI data 393
Figure 5.18 Medway Tunnel Trench 7 CA of all macrofossil data 395
List of Tables
Table 3.1 Plant macrofossil groups and source plant taxa
81
Table 3.2 Taxa and structures inlcuded in the identification research 90-91
Table 3.3 Descriptive characters for rootlets
97
Table 3.4 Descriptive characters for leaves and epidermis
96
Table 3.5 Provisional root types 99
Table 3.6 Summary of leaf and epidermal identification features in the 103
Filicophyta
Table 3.7 Filicophyta trichome types
104
Table 3.8a Summary of the main features of the leaf architecture and 107
epidermis in selected angiosperm taxa - Magnoliflorae and Caryolliflorae
Table 3.8b Summary of the main features of the leaf architecture and
108
epidermis in selected angiosperm taxa - Dilleniflorae and Rosiflorae
Table 3.8c Summary of the main features of the leaf architecture and
109
epidermis in selected angiosperm taxa - Asteriflorae
Table 3.9 Trichome types recorded in the Dicotyledonae
110
Table 3.10 Monocotyledonae
grouped on the basis of leaf architecture and
113
epidermal anatomy
Table 3.11 Main distinguishing features of epidermis and leaves at
114
Class/Order level
Table 3.12 Cover abundance data for trial sample 04/116 119
Table 4.1 Sites included in the project
123
Table 4.2 CANOCO abbreviations for standing vegetation elements
129
Table 4.3 CANOCO abbreviations for macrofossil data
130
Table 4.4 Stonemarsh standing vegetation, sediment and distance data 133
19
Table 4.6 Borstal Marsh standing, vegetation, sediment and distance data 149
Table 4.7 Borstal marsh plant macrofossil data and derived indices
150
Table 4.8 Angel Marsh standing vegetation, sediment and distance data:
162
Transect 1
Table 4.9 Angel Marsh standing vegetation, sediment and distance data:
163
Transect 2
Table 4.10 Angel Marsh plant macrofossil data and derived indices: Transect 167 1
Table 4.11 Angel Marsh plant macrofossil data and derived indices: Block 1
168
Table 4.12 Angel Marsh plant macrofossil data and derived indices: Block 2
169
Table 4.13 Angel Marsh plant macrofossil data and derived indices: Transect
170
2
Table 4.14 Snape Saltings standing vegetation, sediment and distance data
188
Table 4.15 Snape Saltings Transect macrofossil data and derived indices
193
Table 4.16 Snape Saltings area 01 and 03 macrofossil data and derived indices 195
Table 4.17 Snape Saltings Block 1 macrofossil data and derived indices
196
Table 4.18 Snape Saltings Block 3 macrofossil data and derived indices
197
Table 4.19 Snape Saltings: Comparison of seed and fruit data from Blocks 1 204 and 3
Table 4.20 Snape Saltings: Comparison of root cover abundance and 217 corresponding vegetation data
Table 4.21 Burham Marsh East-West Transect and Block sample standing
221
vegetation, sediment and distance data
Table 4.22 Burham Marsh South-North Transect sample standing vegetation, 222
sediment and distance data
Table 4.23 Burham Marsh East-West Transect sample macrofossil data 225
Table 4.24 Burham Marsh South-North Transect sample macrofossil data
226
Table 4.25 Burham Marsh Block I sample macrofossil data
227
Table 4.26 Burham Marsh Block 2 sample macrofossil data 228
Table 4.27 Burham Marsh: Sub-surface sample sediment data from Blocks 1
244
and 3
Table 4.28 Hickling Broad standing vegetation, sediment and distance data
247
Table 4.29a Hickling Broad seed macrofossil data 248
20
Table 4.30 Representation of taxa in the samples from Hickling Broad 261
Table 4.31 Wet Woodland standing vegetation, sediment and distance data
267
Table 4.32a Wet Woodland seed macrofossil data
270
Table 4.32b Wet Woodland non-seed macrofossil data and derived indices 271 Table 4.33 Bure Marsh organic and water data from Block 1 291 Table 4.34 Ubiquity data for taxa identified in the standing vegetation at the 320
sample sites
Table 5.1 Medway Tunnel Section 16 macrofossil records 345
Table 5.2 Medway Tunnel Section 16 main sample groups
361
Table 5.3 Medway Tunnel Section 9 macrofossil records 369
Table 5.4 Medway Tunnel Section 9 sample groups
372
Table 5.5 Medway Tunnel Section 11 macrofossil records
375
Table 5.6 Medway Tunnel Section 11 sample groups 379
Table 5.7 Medway Tunnel Section 6 macrofossil records
383
Table 5.8 Medway Tunnel Section 6 sample groups
388
Table 5.9 Medway Tunnel Trench 7 macrofossil records 392
Table 5.10 Medway Tunnel Trench 7 sample groups
394
Table 5.11 Medway Tunnel (MT93) Phase III charred macrofossil records 398
List of plates
Plate 1 Equisetum fluviatile rootlet with root hairs and epidermis (x100) 552 Plate 2 Equisetum fluviatile rootlet epidermis (x400) 552
Plate 3 Urtica dioica rootlet (x400) 553
Plate 4A triplex portulacoides rootlet epidermis (x 100) 553
Plate 5 Solanum dulcamara rootlet (x100) 554
Plate 6 Lycopus europaeus rootlet (x100) 554
Plate 7 Lycopus europaeus rootlet epidermis (x400)
555
Plate 8 Plantago maritima rootlet epidermis (x400) 555
Plate 9 Galiumpalustris rootlet epidermis (x400) 556
Plate 10 Aster tripolium rootlet (x100)
556
Plate 11 Alisma lanceolata rootlet (x100)
557
Plate 12 Juncus acutorus rootlet (note swollen hair base) (x100)
557
Plate 13 Juncus gerardii rootlet upper and lower epidermal cells (x400) 558
21
Plate 15 Scirpus maritimus rootlet with cubic projections (x100) 559 Plate 16 Glyceria maxima rootlet epidermal cells (x400) 559 Plate 17 Typha latifolia rootlet showing swollen cells (x100) 560 Plate 18 Typha Iatifolia rootlet epidermis (x400) 560 Plate 19 Iris pseudacorus rootlet epidermal detail (x100) 561
Plate 20 Equisetum arvense stem epidermis (x400) 562
Plate 21 Equisetum arvense leaf sheath epidermis (x100) 562
Plate 22 Equisetum variegatum stem epidermis (note silica knobs) (x400)
563
Plate 23 Equisetum variegatum leaf sheath epidermis (x100)
563
Plate 24 Ophioglossum vulgatum venation and epidermis (x40) 564 Plate 25 Ophioglossum vulgatum epidermis detail (x100) 564 Plate 26 Osmunda regalis epidermis with stomata and cells (note the 565
elongated and more regular cells over the vein) (x100)
Plate 27 Polypodium interjectum abaxial epidermis with Type 1 trichome 565 (x 100)
Plate 28 Hymenophyllum tunbrigense pinnule (x40) 566
Plate 29 Hymenophyllum tunbrigense indusium (x40) 566
Plate 30 Pteridium aquilinum marginal false indusium with sporangia (x100) 567 Plate 31 Pteridium aquilinum detail of indusium epidermis (x200) 567 Plate 32 Pteridium aquilinum epidermis with trichomes (x200) 568
Plate 33 Oreopteris limbosperma pinnule (x100) 568
Plate 34 Oreopteris limbosperma epidermis with spherical glands and stomata 569
(x400)
Plate 35 Oreopteris limbosperma type 2 trichome (x 400) 569 Plate 36 Oreopleris limbosperma type 4 trichome (x 400) 570 Plate 37 Thelypteris palustris pinnule showing epidermis and venation (x40) 570 Plate 38 Thelypteris palustris abaxial epidermis (x400) 571
Plate 39Asplenium cetrach adaxial epidermis (x200) 571
Plate 40 Asplenium marinum adaxial epidermis detail (x400) 572
Plate 41 Asplenium marinum abaxial pinnule (x40)
572
Plate 42 Athyrium felix femina epidermis (x400)
573
Plate 43 Polystichum setiferum pinnule apex with spines (x 40) 573 Plate 44 Dryopteris dilitata pinnule apex with spine, epidermis cells and 574
22 Plate 45 Dryopteris dilitata sorus with detail of sporangia and spores (x400) 574
Plate 46 Blechnum spicant epidermis - note thickened lower cells (x100) 575
Plate 47 Anemone nemorosa leaflet margin (x 40) 576
Plate 48 Anemone nemorosa abaxial leaf epidermis with hair (x 100) 576 Plate 49 Anemone nemorosa gland on abaxial leaf epidermis (x 400) 577
Plate 50 Calthapalustris leaf margin with hydathode (x40)
577
Plate 51 Caltha palustris leaf abaxial epidermis (x100)
578
Plate 52 Ramunculus f ammula leaf apex (x40) 578
Plate 53 Ranunculus
flammula abaxial epidermis (x100)
579
Plate 54 Raminculus sceleratus leaflet apex (x40) 579
Plate 55 Ranunculus sceleratus leaflet abaxial epidermis (x400) 580
Plate 56 Ranunculus sub-genus
Batrachium leaflet termination (x100)
580
Plate 57 Glaucium f avum leaf epidermis with hairs (x40)
581
Plate 58 Glaucium
flavum leaf trichome (x100)
581
Plate 59 Glaucium flavum leaf epidermis (x400) 582
Plate 60 Ulmus glabra leaf margin (note cicatrices) (x40) 582
Plate 61 Ulmus glabra leaf vein junction with trichomes (x400)
583
Plate 62 Ulmus glabra abaxial leaf epidermis with trichome (x400) 583
Plate 63 Humulus lupulus trichomes on petiole (x40)
584
Plate 64 Humulus lupulus abaxial leaf epidermis (x100)
584
Plate 65 Urtica dioica leaf apex (x40) 585
Plate 66 Urtica dioica abaxial leaf epidermis (x100) 585
Plate 67 Urtica dioica type 3a trichome (x100)
586
Plate 68 Myrica gale abaxial leaf epidermis (x100)
586
Plate 69 Myrica gale adaxial leaf epidermis (x100)
587
Plate 70 Myrica gale leaf edge (x100)
587
Plate 71 Fagus sylvatica leaf edge (x40) 588
Plate 72 Fagus sylvatica leaf lamina (x40)
588
Plate 73 Fagus sylvatica abaxial leaf epidermis (x400)
589
Plate 74 Quercuspetraea apex of leaf lobe (x40)
589
Plate 75 Quercuspetraea type 14 and 15 trichomes on abaxial leaf epidermis
590
(x400)
Plate 76 Quercus petraea type 16 trichome on abaxial leaf epidermis (x400) 590
23
Plate 78 Quercus robur abaxial leaf epidermis (x400) 591
Plate 79 Alnus glutinosa leaf edge (x40) 592
Plate 80 Alnus ghrtinosa hairs in vein axil on abaxial leaf surface (x40)
592
Plate 81 Alnus glutinosa leaf showing areolation (x40) 593 Plate 82 Almis glutinosa abaxial leaf epidermis (x400) 593
Plate 83 Betulapendula leaf margin with hairs and glands (x40)
594
Plate 84 Betulapendula trichomes on primary leaf vein (x40) 594
Plate 85 Betula pendula abaxial leaf epidermis and crystals in bundle sheath
595
(x400)
Plate 86 Betula pendula peltate gland on leaf epidermis in plan view (x400) 595 Plate 87 Carpinus betulus leaf marginal tooth (x40) 596 Plate 88 Carpinus betulus leaf mesophyll crystals under polarised light (x100) 596 Plate 89 Corylus avellana abaxial leaf epidermis (x400) 597
Plate 90 Corylus avellana abaxial leaf epidermis with trichomes (x100)
597
Plate 91 Corylus avellana abaxial epidermis with trichome type 2 (x100) 598 Plate 92 Atriplex portulacoides leaf margin with detached spherical glands 598
(x100)
Plate 93 Atriplex portulacoides leaf abaxial epidermis (x400) 599 Plate 94 Atriplex portulacoides glands above leaf epidermis (x100) 599
Plate 95 Salicornia sp. epidermis (x400) 600
Plate 96 Sarcocorniaperennis stem epidermis (x100) 600
Plate 97 Suaeda maritima leaf epidermis (x100) 601
Plate 98 Cerastiumfontana abaxial leaf epidermis (x100)
601
Plate 99 Cerastiumfontanum stomata on abaxial leaf epidermis (x400) 602
Plate 100 Persicaria hydropiper leaf edge (x100) 602
Plate 101 Persicaria hydropiper leaf edge (x40) 603
Plate 102 Persicaria maculosa abaxial leaf epidermis (x400) 603
Plate 103 Rumex acelosella gland on abaxial surface (x400)
604
Plate 104 Persicaria hydropiper leaf edge with trichome type 19 (x100) 604
Plate 105 Rumex hydrolapathum leaf edge (x40)
605
Plate 106 Rumex hydrolapathum abaxial leaf epidermis (x100)
605
Plate 107 Armeria maritima leaf epidermis (x100) 606
Plate 108 Armeria maritima leaf epidermis (x400) 606
24
Plate 110 Limonium sp. abaxial leaf epidermis showing salt gland and stomata 607 (x400)
Plate 111 Limonium sp. leaf edge (x40)
608
Plate 112 Tilia cordata leaf margin (x40) 608
Plate 113 Tilia cordata leaf venation and areolation (x40) 609 Plate 114 Tilia cordata abaxial leaf epidermis (note striations) (x400) 609 Plate 115 Althaea officinalis leaf margin showing stellate trichomes (x40) 610
Plate 116 Althaea officinalis trichomes under polarised light (x40)
610
Plate 117 Althaea officinalis abaxial leaf epidermis (x400)
611
Plate 118 Viola sp. leaf edge (x400) 611
Plate 119 Viola sp. abaxial epidermis (x400) 612
Plate 120 Salix cinerea leaf margin with apical gland (x40) 612 Plate 121 Populus nigra abaxial leaf epidermis (x100) 613
Plate 122 Populus tremula leaf edge (x40) 613
Plate 123 Salix cinerea abaxial leaf epidermis (x400 614 Plate 124 Rorippa sp. venation on leaf under polarised light (x40) 614
Plate 125 Rorippa sp. abaxial leaf epidermis (x100)
615
Plate 126 Cochlearia anglica abaxial leaf epidermis (x400) 615 Plate 127 Lysimachia vulgaris abaxial leaf epidermis (x100) 616
Plate 128 Lysimachia vulgaris abaxial leaf epidermis (x100)
616
Plate 129 Crataegus monogyna leaf edge (x40) 617
Plate 130 Filipendula ulmaria leaf serration (x40) 617
Plate 131 Filipendula ulmaria abaxial epidermis (x100)
618
Plate 132 Filipendula tdmaria leaf edge and venation (x400) 618 Plate 133 Geum rivale leaf vein with type 22 trichome (x100) 619
Plate 134 Prunus spinosa leaf edge (x100)
619
Plate 135 Prunus spinosa abaxial epidermis (x400) 620
Plate 136 Rosa sp. leaf serration (x40) 620
Plate 137 Rosa sp. adaxial leaf epidermis (x100)
621
Plate 138 Rosa sp. abaxial epidermis (x100)
621
Plate 139 Rosa sp. trichomes on leaf vein (x400)
622
Plate 140 Rubus fruticosus leaf edge (x40) 622
25
Plate 142 Sorbus aucuparia serration apex (x40) 623
Plate 143 Lathyruspalustris leaf edge (x40) 624
Plate 144 Lathyruspalustris abaxial leaf epidermis (x400) 624
Plate 145 Lythrum salicaria leaf edge (x40) 625
Plate 146 Epilobium hirsutum leaf edge (x40)
625
Plate 147 Epilobium hirsutum raphides in leaf mesophyll (x100)
626
Plate 148 Epilobium hirsutum abaxial leaf epidermis (x400) 626
Plate 149 Cornus sanguinea adaxial leaf epidermis with trichome (x100)
627
Plate 150 Cornus sanguinea abaxial epidermis (x400) 627
Plate 151 Rhamnus cathartica leaf margin (x40) 628
Plate 152 Rhammis cathartica abaxial leaf epidermis (x400) 628
Plate 153 Acer campestre leaf margin (x40)
629
Plate 154 Acer campestre abaxial leaf epidermis (x100) 629 Plate 155 Acer campestre peltate gland on abaxial leaf epidermis (x400) 630
Plate 156 Hedera helix leaf edge (x40) 630
Plate 157 Hedera helix abaxial leaf epidermis (x100) 631
Plate 158 Apium graveolens leaf margin (x40) 631
Plate 159 Apium graveolens abaxial epidermis (x400) 632
Plate 160 Apium nodorum leaf edge (x40) 632
Plate 161 Calystegia sepium leaf edge (x 40) 633
Plate 162 Calystegia sepium abaxial epidermis (x100) 633 Plate 163 Calystegia sepium adaxial leaf epidermis (x100) 634
Plate 164 Myosotis scorpioides leaf edge (x40) 634
Plate 165 Myosotis scorpioides leaf edge detail (x100) 635
Plate 166 Myosotis scorpioides abaxial epidermis (x40)
635
Plate 167 Veronica beccabunga leaf edge (x40) 636
Plate 168 Veronica beccabunga abaxial leaf epidermis (x400) 636
Plate 169 Lamium album leaf margin (x100) 637
Plate 170 Lamium album abaxial leaf epidermis cicatrice (x100) 637
26
Plate 174 Lycopus europaeus hair detail (x400) 639
Plate 175 Plantago coronopus leaf lobe (x40) 640
Plate 176 Plantago coronopus leaf epidermis (x400) 640 Plate 177 Plantago maritima leaf epidermis (x400) 641
Plate 178 Fraxinus excelsior leaf margin (x40) 641
Plate 179 Fraxinus excelsior abaxial leaf epidermis with peltate gland (x400) 642 Plate 180 Scrophularia nodosa trichomes on abaxial leaf surface (x100) 642 Plate 181 Galium pahutre leaf margin (note raphides) (x40) 643
Plate 182 Galium palustre adaxial leaf epidermis (x100)
643
Plate 183 Gall um palustre abaxial leaf epidermis (x400) 644 Plate 184 Lonicera periclymenum leaf margin (x100) 644
Plate 185 Sambucus nigra leaf margin (x40) 645
Plate 186 Sambucus
nigra abaxial leaf epidermis (x100)
645
Plate 187 Sambucus
nigra abaxial leaf epidermis (x400)
646
Plate 188 Viburnum opulus leaf tip (x40) 646
Plate 189 Viburnum opulus abaxial leaf epidermis (x400)
647
Plate 190 Bidens cenma leaf margin (x40) 647
Plate 191 Bidens cernua abaxial epidermis (x400) 648
Plate 192 Cirsium palustre leaf margin with trichome (x40) 648
Plate 193 Cirsium pahistre abaxial leaf epidermis (x400)
649
Plate 194 Eupatorium cannabinnim leaf margin (x40) 649 Plate 195 Eupatorium cannabinum trichomes on leaf vein (x100) 650
Plate 196 Eupatorium cannabintim abaxial leaf epidermis with gland (x400)
650
Plate 197 Butomus umbellatus leaf epidermis (x100) 651
Plate 198 Alisma lanceolata leaf epidermis (x100)
651
Plate 199 Sagittaria sagittifolia leaf epidermis (x100) 652
Plate 200 Hydrocharis morsus-ranae leaf epidermis with stomata and
652
mesophyll air-spaces (x100)
Plate 201 Hydrocharis morsus-ranae leaf epidermis (x400) 653
Plate 202 Potamogeton lucens leaf margin (x100)
653
Plate 203 Juncus acutus stem epidermis (x100)
654
Plate 204 Juncus maritimus stem epidermis (x100) 654
Plate 205 Carex remota leaf epidermis with stomata (x400) 655
27
Plate 207 Eleocharispalustris leaf edge (x400)
656Plate 208 Scirpus lacustris stem epidermis with stomata and prickle hair 656 (x400)
Plate 209 Scirpus lacustris leaf epidermis with stomatal files (x400) 657 Plate 210 Scirpus maritimus stem epidermis with stomata and hairs (x400) 657 Plate 211 Elymus sp. leaf epidermis showing long and short cells (x400) 658 Plate 212 Phalaris arundinacea leaf epidermis (x 400) 658 Plate 213 Phalaris arundinacea stem epidermis showing spines (x400) 659 Plate 214 Typha latifolia leaf epidermis showing stomatal files (x400) 659
28
1 Introduction
Alluvium has been deposited in many rivers and estuaries in the Lower Thames Basin (referred to from here as the `Lower Thames') during the Holocene. As river valleys and
estuaries are the only spatially extensive sedimentary basins in southeast England in which long sedimentary sequences have been deposited, alluvium is a particularly
important source of data for regional archaeological and palaeoenvironmental studies. Alluvium has provided data for the study of Holocene environmental patterns and
processes at micro-, macro- and mega-spatial scales in the region (sensit Delcourt et al. 1983), including terrestrial ecological change (Scaife and Burrin 1987; Greig 1992a,
1992b; Thomas 1996), Holocene river hydrology (Burrin and Scaife 1984) and relative sea-level change (Devoy 1979). Archaeological research over 150 years has demonstrated that alluvial environments were the site of direct human activity throughout the Holocene, with finds including settlements, boats, artefacts, trackways
and platforms (see review of Bates and Barham 1995). They also preserve important traces of life in adjacent terrestrial locations that for millennia have been important
centres of population, industry and trade. High sediment accumulation rates and maintained anaerobic conditions have led to the preservation of organic archaeological materials that would otherwise perish.
Interpretation of human activity and reconstruction of patterns of environmental change in and adjacent to alluvial environments has best been achieved through the
systematic integration of palaeoenvironmental and archaeological data. This approach has a long history (e. g. Bulleid and Gray 1911,1917; Reid 1917, Clifford 1936, Swinnerton 1931, Godwin 1960) and has been an explicit part of several recent research programmes in wholly or partially alluviated areas (Coles 1987; Van de Noort and Davies 1993; Cowell and Inns 1994; Wilkinson and Murphy 1995). Such systematic
programmes have been largely absent from the Lower Thames, with the exception of the Blackwater Estuary Project, and most studies have been associated with individual
rescue archaeology works, the reports from which are dispersed in unpublished archives or obscure local publications.
Although alluvium is a regionally important source of information, its use as a
basis for reconstructions of environments
in time and space is complicated by its mode of
29
inputs from both allochthonous and autochthonous sources. Deposition is also often discontinuous and may vary in rate and direction, with sediments subjected to episodes of erosion. Sediments may, therefore, contain widely varying concentrations of fossils differing with spatial and temporal fidelities (sensor Kidwell and Flessa 1995) and
sequences may be incomplete, with important transitional sections missing. This has partially led to the restricted use of alluvial fossil assemblages for palaeoenvironmental investigations, especially those from clastic sediments. A further problem is that alluvial environments are spatially heterogenous with many sub-environments that have differing
sedimentological and fossil characteristics existing at the same point in time. Changes in sediments and fossils may simply reflect local changes and the linkage of fades changes to larger environmental variables is often difficult to prove.
Among the fossil groups preserved in alluvium are plant macrofossils. This group
includes all plant structures visible to the naked eye, such as seeds, wood, roots, rhizomes, stem and leaf fragments (Dickson 1970; Birks and Birks 1980). Macrofossil preservation in alluvium is often excellent owing to the anaerobic environment of many
sediments. However, analysis of plant macrofossil assemblages is one of the more neglected areas of alluvial research. Descriptions are often limited to the partial identification of fibres, stems and leaves visible in superficial examination of strata (e. g. Waller 1994) or the recording of gross macrofossil groups in stratigraphie descriptions such as those based on the Troels-Smith system (Aaby and Berglund 1986). These records primarily contribute to the identification of depositional environments and, in some cases, the type of vegetation extant during sediment accumulation. Mostly, only loose structural vegetation groups are referred to (e. g. carr, reedbeds, sedge beds) which may be of limited use for investigating detailed ecological variables and environmental
parameters. As in most areas of palaeoenvironmentäl study, macrofossil analysis has become an adjunct to palynology when detailed vegetation reconstructions are required.
When used, completed macrofossil analyses are usually restricted to the identification and quantification of a limited range of macrofossils, usually only seeds and fruits. Many
classes of remains are not included in analysis or are subject to only cursory examination and rarely if ever subject to quantification and quantitative analysis.
Macrofossils have the potential to make a greater contribution to the
investigation of alluvial facies than has previously been the case by allowing greater
30
source for spatially precise vegetation information, useful for detailed palaeoecological investigations and contextualising archaeological finds. This potential is due to the heterogeneity of macrofossil assemblages, the limited dispersability of many plant parts in
sedimentary environments and the potentially high level of taxonomic identification.
Macrofossils in general are deposited close to the site of plant growth (Collinson 1983; Field 1992) and therefore provide spatially precise information about former vegetation. Such information is important if detailed palaeoecological data are to be
gained and if the physical context of archaeological sites is to be understood, a vital element in any interpretation. Alluvial ecosystems have been modified or destroyed by human activity in the last five centuries and palaeoecological information may be the only way to investigate many elements of `natural' alluvial ecosystems. Macrofossil analysis
also provides the potential for much more detailed identifications of ecosystem elements as many macrofossils can be identified to genus and species level, unlike many pollen types.
Identification of vegetation types at a sample point and identification of local
vegetation responses is important in determining local and more widespread changes in environmental conditions. Macrofossil assemblage analysis also has the potential to contribute to the identification of the depositional environment. Macrofossil assemblages
consist of groups of plant structures (e. g. roots, stems, leaves) and tissues (e. g. cuticles, wood) as well as taxa and, therefore, may be described in structural as well as taxonomic
terms. These properties are potentially useful in differentiating depositional environments when differentiation through other means may be impossible, a common problem in
alluvial sequences in the Lower Thames.
Although a potentially useful source of information, detailed macrofossil analysis
31
problem in macrofossil analysis of, alluvium is access for sampling. Extensive exposures
of sediments
are rare in many areas of southeast England and it is still unclear whether
coring and monolith sampling is adequate for macrofossil collection and the sample sizes
that are adequate
for different levels of macrofossil-based
information.
The objective of the research presented in this thesis is to develop, test and
evaluate analytical approaches to plant macrofossils from alluvial facies, specifically
targeted at the southeast region of England. It aims to:
I.
Develop recording and identification techniques for analysis of macrofossil
assemblages,
establishing
identification criteria where they are lacking;
II.
Investigate plant macrofossil assemblage
formation processes through actualistic
research
(sensu Kidwell 1986) in modern alluvial environments to determine:
A.
the potential of different alluvial depositional sub-environments to
incorporate plant macrofossils;
B. the quantitative and qualitative properties of macrofossil assemblages preserved in alluvial depositional sub-environments;
C.
the spatial and temporal resolution of information about vegetation
structure and floristics provided by different classes of macrofossils and
whole macrofossil assemblages
preserved in different alluvial depositional
sub-environments;
D. the effects of sample size on the information attainable through macrofossil analysis in different alluvial depositional sub-environments.
III.
Evaluate the approach developed through application of an integrated analysis to
macrofossil assemblages
recovered from a Holocene alluvial sequence
in which
palaeoenvironmental
and archaeological
material is preserved.
The following chapters provide background information to the research, details of the
methods, sites used and results obtained, and also evaluates these results. Chapter 2
considers the value of alluvium as a resource for archaeology and palaeoecology in the
Lower Thames and includes a brief consideration of rivers and estuaries as sedimentary
32
Chapter 3 presents the methods used to generate macrofossil data. It includes a classification of macrofossils, criteria for the identification of previously difficult macrofossil groups and a discussion of the method of macrofossil quantification used in the subsequent chapters. Much of the work in this chapter aims to develop a usable
analytical method and the broad range of subjects covered by it means that some had to be treated in a cursory manner.
Chapter 4 details observations of macrofossil accumulation and representation in
a range of modern depositional environments including tidal mudflats, saltmarshes and freshwater swamps, reedbeds, herb fens and wet woodlands. Data collected from these observations have provided a basis for discussing macrofossil taphonomy and evaluating the spatial and temporal resolution of information derived from macrofossil assemblages in different depositional environments. The focus was mainly on non-seed macrofossil
assemblages, although seed concentrations and profiles were investigated for most of the sites. The observations are also used to determine the suitability of different macrofossil categories and groups of macrofossils to investigate ecological and environmental parameters such as salinity gradients, vegetation structure and vegetation floristics.
Chapter 5 presents the analysis of a group of plant macrofossil assemblages
from
sediments sampled during the construction of the Medway Tunnel in Chatham, Kent. Sediment accumulation was initiated at ca 7000 BP at the site and subsequent alluviation preserved Mesolithic, Neolithic, Bronze Age, Iron Age and Romano-British
archaeological features and artefacts within a complex series of alluvial facies. Plant macrofossil analysis aimed to contribute to the definition of depositional environments, reconstruction of local and regional palaeoenvironments and act as a basis for evaluating the efficacy of the developed macrofossil methodology.
Chapter 6 discusses
and evaluates the research. Deficiencies and strengths of the
33
2 Plant macrofossil analysis in archaeological and palacoenviron mental
investigations of Holocene alluvium, with special reference to the Lower Thames Basin
2.1 Nature and distribution of Holocene alluvium
Alluvium is sediment deposited along river courses in fluvial, estuarine and terrestrial environments (Bates and Barham 1995). Thick deposits accumulated during the Holocene in the rivers of the Lower Thames Basin grading into coastal sediments. The River Thames is the major river of the region which, with its tributaries, drains the Thames Basin, sections of the surrounding Downs and the north Weald (Figure 4.1). To the north of the Thames lie the southern Essex rivers of the Colne, Crouch and Blackwater and to the southeast the River Medway, the main northern conduit of water
from the Kentish Weald.
In this text the definition is used to denote all of the sediments that may be
deposited in the diverse environments of river-estuarine systems. The following discussion also refers to sites that would usually be considered as `coastal' as opposed to riverine in genesis. This reflects the blurred boundaries between sedimentation processes
and outcomes in these environments. It also reflects the complexity of Holocene alluvial sediment sequences that contain riverine, estuarine and coastal sediments and, therefore, the wide range of methods that their analysis requires. Many of the sites can be considered within alluvial archaeology and inter-tidal archaeology. The following has been derived from numerous sources, but draws heavily on Reading (1978), Dyer (1979), Reineck and Singh (1980), Lewin (1981), Richards (1982), Pethick (1984),
Collinson (1986), and Allen (2000).
Holocene sediments have been grouped together on the Thames as the Tilbury
Formation (Bridgland 1995), but this grouping belies an inherent complexity. The Tilbury Formation includes a great variety of sediments dominated by inter-tidal muds and organic rich muds and peats, forming a characteristic alternating pattern of peat bands and inorganic units in several rivers. The number and thickness of peat beds varies with location, up to five being identified at Tilbury (Devoy 1979,1982). Holocene sediments fill the inherited Pleistocene landscape to a height of approximately 5m
ordnance datum (OD) reaching depths, depending on the underlying topography, of between
34
north Kent Marshes and Medway floodplains (Skempton 1995; Bates et al. 1995). The areal extent of alluviated areas is considerable in the region, forming a broad band along
the Thames of up to 4.5km wide.
2.2 Rivers and estuaries as sedimentary basins
Rivers are the major agents of terrestrial erosion and transportation, carrying water and sediment from the land to the sea. They form a continuum with estuaries in which freshwater discharge mixes with saline water flowing into a river mouth due to tidal action. Rivers and estuaries are connected but the environments of each are dominated by different hydrological and sedimentation processes. They also contain a range of
depositional and erosional environments in which different landforms are generated, inhabited by a diverse biota and represented in the stratigraphic record by a wide range of sedimentary facies.
2.2.1 Hydrological factors
Rivers form as the result of the flow of excess water into channels that conduct water to the sea and entrain, transport and deposit sediment. The main upstream controls on
water supply, sediment supply and catchment are the basin physiography, climate, geology, vegetation, soils and land-use. Base-level, the level to which water flows as the result of gravity, sets the downstream control and is set locally by the presence of erosion-resistant rocks and ultimately by the sea at the river-mouth (Leopold et at
1964).
The initial quantity of water available to a river is a balance between precipitation
and evapotranspiration (determined by climate) and the capacity of soils and rocks to
absorb the water (Richards 1982) enhanced
by the presence of vegetation. Water enters
into rivers as the result of
a) Infiltration excess overland flow, when precipitation exceeds the soil's capacity to
absorb it;
35
Discharge, the quantity of water passing a given point in a river channel, varies over the year, usually in a regular cycle. Porous base rocks, such as chalk in the Lower Thames, can absorb rainfall and release it slowly, decreasing peak discharge (Ward
1981). Water velocity is a function of discharge, basin-slope, gravity and the composition of the riverbank, an important factor in determining the resistance of the riverbanks to erosion and channel transformation. Floodplain water levels are affected by the periodicity of flooding, groundwater levels, precipitation and permeability of the substrate, perched water tables forming where clay-silt substrates impede groundwater drainage (Brown 1997).
Estuarine hydrology is dominated by the diurnal tidal cycle. Rising tides cause
water to be forced up river valleys, the distance being determined by tidal range, basin
shape and river discharge.
Estuaries in southeast England are macro-tidal, having a tidal
range of 4m, and tidal incursion dominates river discharge. Maximum velocities are
reached at mid-tide, minimum velocities being reached at high and low tide. Tides
influence landforms through direct inundation and the ponding of river discharge behind
the tidal wave. Wind waves are relatively unimportant except in storms, when, like
secondary
currents caused by mixing of fresh and saline water, they can introduce marine
sediments
to the system.
2.2.2 Sedimentation
