Chesapeake Bay Living Resources Task Force
QH5415E8H31
QH
541.5
.E8
H32
1987
August 1987
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HABITAT REQUIREMENTS
FOR CHESAPEAKE BAY LIVING RESOURCES:
A Report from the Chesapeake Bay
Living Resources Task Force
Annapolis, Maryland
August, 1987
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DISCLAIMER
This report has been reviewed by the Living Resources Task Force of the Chesapeake
Bay Implementation Committee and approved for publication by the Chesapeake Bay
Program, U.S. Environmental Protection Agency. Approval does not signify that the
contents necessarily reflect the view and policies of the U.S. Environmental Protec-
tion Agency, nor does mention of trade names or commercial products constitute
endorsement or recommendation for use.
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ACKNOWLEDGEMENTS
The Chairman of the Living Resources Task Force would like to acknowledge the par-
ticipation and contributions of: the members and supporting staff of the Chesapeake
Bay Living Resource Task Force; participants in the February Workshop on Habitat
Requirements for Chesapeake Living Resources; the principal authors of the report,
Steve Jordan, David Pyoas, and Charles Frisbee of the Maryland Department of
Natural Resources and Bert Brun of U.S. Fish and Wildlife; the technical editor, Nina
Fisher, Chesapeake Bay Program/Computer Sciences Corporation; and, the scientific
editor, Bess Gillelan, Chesapeake Bay Program/Computer Sciences Corporation.
MEMBERS OF THE CHESAPEAKE BAY LIVING RESOURCES TASK FORCE
Ralph Abele
Pennsylvania Fish Commission
Louis Bercheni
Pennsylvania Department of
Environmental Resources
Glenn Kinser
U.S. Fish and Wildlife Service
Louis Sage
Academy of Natural Sciences
Charles Spooner
U.S. EPA Chesapeake Bay Program
Elizabeth Bauereis
Baltimore Gas and Electric Company
Steve Jordan
Maryland Department of
Natural Resources
Larry Minock
Virginia Council on the Environment
Robert Siegfried
Virginia Water Control Board
James Thomas
NOAA Estuarine Programs Office
Lee Zeni
Interstate Commission on the
Potomac River Basin
KEY
m = meter
C = celcius
ppt = parts per thousand
KD = light attenuation coefficient
TRC = total residual chlorine
cm/s = centimeters per second
chlor. = chlorophyll
mg/1 = milligrams per liter - equivalent to parts per million
ug/1 = micrograms per liter - equivalent to parts per billion
LCO = lethal concentration - 0 percent mortality
LC50 = lethal concentration - 50 percent mortality
urn = micron
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FOREWORD
The Living Resources Task Force, an ad hoc workgroup of the Chesapeake Bay
Program, was charged by the Chesapeake Bay Implementation Committee to develop
an approach to define habitat objectives for the living resources of the Bay. The
objective of the Task Force in producing this report was to establish a technically
defensible approach in setting regional habitat objectives for Chesapeake Bay by
initially assembling habitat requirements for individual target species. The scope of
this report places limitations on its utility as a planning document for Bay managers.
It is intended, however, to summarize the results of the Task Force efforts to date and
to provide the basis for future refinement of the habitat objectives approach. This
document describes the results of ongoing efforts to identify critical habitat require-
ments for target species.
Within the context of this report, habitat is defined as the biotic and abiotic con-
ditions upon which the living resources of the Bay depend. Abiotic conditions
include factors such as water quality, substrate, circulation patterns, bathymetry,
and weather; two dominant factors are salinity and depth. Biotic conditions are
governed by variables such as vegetative cover, quality and quantity of prey species,
species composition, population density, and primary productivity. The estuarine
environment represents a wide range of these conditions which are dynamic in time
and space. Although Bay species are tolerant of dynamic natural conditions, their
habitats have been altered by man-induced activities; there is evidence that
thresholds for tolerating adverse conditions have been exceeded. The Living
Resources Task Force has attempted to identify the boundaries of tolerable conditions
in the form of habitat requirements.
The report is constructed following the guidelines created to direct the develop-
ment of living resources habitat requirements. The sections on the Chesapeake Bay
ecosystem and the major physical factors affecting the Bay provide the structural
framework for all subsequent discussions of the living resources. The representative
living resources are a group of organisms that serve as indicators of the Bay's
ecological condition. From this group, target species were selected as particularly
important for the development of initial habitat requirements. The report includes a
set of matrices outlining habitat requirements for critical life stages of the target
species as well as range maps of these stages.
A scientific workshop, with invited participants from universities, research in-
stitutions, and state and federal agencies, was held to review the initial list of
requirements and advise the Living Resources Task Force on critical life stages of the
target species and seasonal and geographic distributions of the critical life stages.
The workshop proceedings are contained in Appendix C: Report of the Workshop on
Habitat Requirements for Chesapeake Bay Living Resources (Connery, 1987).
To guide subsequent efforts in linking living resources to habitat conditions,
several recommendations for future tasks are proposed. These include expanding the
habitat matrices to encompass requirements for food species on which the target
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species depend, creating habitat matrices for other representative species, identi-
fying species and population characteristics that could serve as indicators of the
Bay's health, and encouraging Bay planners to incorporate habitat requirements
into their environmental planning efforts.
This report will be utilized during discussions leading to the signing of the
revised Chesapeake Bay Agreement in December 1987. Continued development of
habitat and living resource goals will be part of the focus in the implementation of
that Agreement.
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TABLE OF CONTENTS
ACKNOWLEDGEMENTS iii
FOREWORD v
I. INTRODUCTION 1
II. THE CHESAPEAKE BAY ECOSYSTEM 5
A. Plankton 5
B. Vegetation 7
C. Benthos 9
D. Finfish 9
E. Waterfowl and Wildlife 10
III. CHESAPEAKE BAY HABITAT ZONATION 11
A. Depth Zones 11
B. Salinity Zones 12
IV. SPECIES SELECTION 15
A. Representative Species 15
B. Target Species 19
V. HABITAT MATRICES 21
Target Species: Submerged aquatic vegetation complex 21
Target Species: Striped bass (Morone saxatilis) 24
Target Species: Alewife (Alosa pseudoharengus) and
blueback herring (Alosa aestivalis) 26
Target Species: American shad (Alosa sapidissima) and
hickory shad (Alosa mediocris) 30
Target Species: Yellow perch (Perca flavescens) 33
Target Species: White perch (Morone americana) 35
Target Species: Menhaden (Brevoortia tyrannus) 38
Target Species: Spot (Leiostomus xanthurus) 38
Target Species: Bay anchovy (Anchoa mitchilli) 41
Target Species: Molluscan shellfish: American oyster
(Crassostrea virginica), soft clam (Mya arenaria)
and hard clam (Mercenaria mercenaria) 43
Target Species: Blue crab (Callinectes sapidus) 49
Target Species: Canvasback (Aythya valisineria) 51
Target Species: Redhead duck (Aythya americana) 51
Target Species: Black duck (Anas rubripes) 56
Target Species: Wood duck (Aix sponsa) 56
Target Species: Great blue heron (Ardea herodeas) 60
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Target Species: Great (American) egret (Casmerodius albus) 60
Target Species: Little blue heron (Florida caerulea) 63
Target Species: Green heron (Butorides striatus) 63
Target Species: Snowy egret (Egretta thula) 63
Target Species: Bald eagle (Haleaeetus leucocephalus) 67
Target Species: Osprey (Pandion halaetus) 67
VI. LITERATURE CITED 72
VII. SELECTED REFERENCES 79
References for Representative Species of Finfish Cited in Chesapeake Bay
Habitat Matrices 83
References for Representative Species of Shellfish Cited in Chesapeake Bay
Habitat Matrices 84
References for Representative Species of Birds Cited in Chesapeake Bay
Habitat Matrices 84
APPENDIX A: TOXICITY OF SUBSTANCES TO STRIPED BASS LARVAE AND
JUVENILES - ADAPTED FROM WESTIN AND ROGERS, 1978
APPENDIX B: HABITAT DISTRIBUTION MAPS FOR THE CRITICAL LIFE STAGES
OF THE TARGET CHESAPEAKE BAY LIVING RESOURCE SPECIES
APPENDIX C: REPORT OF THE WORKSHOP ON HABITAT REQUIREMENTS FOR
CHESAPEAKE BAY LIVING RESOURCES
Vlll
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INTRODUCTION
Declines in stocks of finfish, shellfish, waterfowl and submerged aquatic
vegetation in the Chesapeake Bay have prompted an unprecedented effort by
the states and federal government to understand causes of the declines and to
explore means of restoring and protecting these stocks. Studies completed in
1983 under the aegis of the Environmental Protection Agency concluded that
the decline of important resources was due to deteriorating water quality, par-
ticularly nutrient enrichment and contamination by toxic metals and organic
compounds (EPA, 1983).
Since 1983, most of the research and planning efforts for restoring and
protecting the Chesapeake Bay has focused on documenting the present water
quality of the Bay and refining strategies for reducing or preventing further
increases in nutrient and contaminant loads. Strategies based primarily upon
water quality, however, cannot necessarily ensure the restoration and pro-
tection of living resources. The most tangible warning signs of widespread
environmental problems in the Bay have been shifts in the relative abun-
dance of living resources. Therefore, living resources serve as excellent indi-
cators of the Bay's recovery for Bay managers and the public.
The abundance and distribution of species within the Bay are related to
many variables: climate, natural population cycles, reproductive potential,
disease, predation, and the abundance and quality of food and habitat. Human
activities impose another set of conditions which both directly and indirectly
affect species abundance. Fishing, land and water uses, contaminant dis-
charges, and physical habitat alterations can directly affect important species.
Indirect impacts of these activities can result in disruption of food chains and
perturbation of the ecological balance of the estuary.
In recognition of these principles, the Chesapeake Bay Program's
Implementation Committee established the Living Resources Task Force (LRTF)
to develop a living resource-based approach for defining habitat objectives
for the Bay. The membership of the LRTF consisted of managers and scientists
from federal and state agencies, private industry, and universities. Through a
series of meetings at both the managerial and technical levels, the Task Force
outlined an approach to establish living resource objectives by first identify-
ing habitat requirements for selected target species. The habitat requirements
are intended to provide planners, managers, researchers, and modelers of the
Bay with information on the minimum habitat quality needed by the target
species and the plants and animals upon which the target species depend for
food. These requirements can be used to estimate the feasibility, benefits and
potential costs of maintaining and protecting an estuarine environment
suitable for the successful reproduction and survival of living resources.
Habitat requirements are not meant to be standards or criteria for wastewater
discharge permitting or other types of regulatory activities, but they can be
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used to develop water quality standards for regions of the Bay that are defined
in terms of living resource habitat rather than water use.
The relationship between the restoration or protection of living
resources and requirements for protecting specified habitats requires clarifi-
cation. Achievement of the proposed requirements will not necessarily
directly result in the establishment of specific population or harvest levels for
any of the targeted species. For example, total compliance with requirements
for striped bass larvae may not result in an improvement of the annual
juvenile index. However, the recovery of species which have declined in
Chesapeake Bay and the reestablishment of a balanced ecosystem must be seen
as the ultimate measures of success in restoring the quality of Chesapeake Bay.
These goals will be unattainable unless certain minimum habitat requirements
are achieved.
The Living Resources Task Force used the following sequential guidelines
for developing the living resources habitat requirements described in this
document:
1. Representative species for the Chesapeake Bay
were identified for all trophic levels, including
plankton, vegetation, benthic organisms, shellfish,
finfish, and wildlife;
2. A smaller group of target species were identified
for immediate development of habitat requirements.
Criteria selecting the target species were based upon
their commercial, recreational, aesthetic, or ecological
significance and the threat to sustained production due
to population decline or serious habitat degradation;
3. The critical life stages and critical life periods
for the target species were identified;
4. Habitat requirement matrices for the targetted
living resources and the species upon which they
prey were developed and refined from current scientific
literature and recent research findings;
5. Geographic areas of the Bay were defined where
habitat requirements should be met in order to protect
the reproduction and survival of the target species. These
areas were based upon present distributions with
consideration also given to historical distributions.
The guidelines were not set up to address issues of numerical population
objectives or management of fish and game harvests. For most Chesapeake Bay
species, neither the total population size nor the information needed to esti-
mate stock sizes is available at present, so realistic objectives for population
sizes cannot be set. While meeting habitat criteria may not ensure survival of
a species in the face of exploitation, there can be no harvest in the absence of
sufficient suitable habitat to support the species. The purpose of this first
phase of the Task Force effort is to specify the quality and geographic distri-
bution of Bay habitats necessary for the sustainable reproduction and long-
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term survival of the target species. In the future, the living resources
restoration efforts may also address such issues as:
1. Establishment of additional habitat requirements
that support both prey of the target species and
other representative species. Special attention should
be paid to the planktonic and benthic communities as
indicators of ecosystem stress and as support organisms
for higher trophic levels;
2. Identification of those characteristics of living resource
populations (e.g. distribution and abundance) or of Bay
communities (e.g. diversity) that will serve as
measures of the Bay's recovery or lack of recovery
in response to management actions;
3. Provisions for refining programs for monitoring, living
resources and habitat conditions, as well as water quality,
and for using computer models of the Bay to predict
the effects of actions to improve habitat conditions,
such as nutrient reduction strategies;
4. Synthesis of habitat requirements into regional habitat
objectives.
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THE CHESAPEAKE BAY ECOSYSTEM
Public interest in the environment has centered directly on the
Chesapeake Bay's aesthetic and economic values and indirectly on its eco-
logical values. The success of economically-important finfish and shellfish is
ultimately dependent on the primary producers of the Bay -- phytoplankton
and other organisms that form the base of the Chesapeake's food chain. The
animals, plants, and microbes of the Bay are interwoven by a complex of
feeding, chemical, and physical interactions. Thus, successful restoration and
protection of commercially, recreationally, and ecologically-important species
are not solely dependent upon the physical and chemical integrity of habitats:
the integrity of the trophic food web supporting these populations is crucial to
resource survival and abundance.
Figure 1 is a network diagram of the summer, mesohaline Chesapeake Bay
designed by Ulanowicz and Baird (1986). The network is presented as a proto-
type of the major trophic relationships and energy pathways in the Bay. It
has been greatly simplified (in comparison to the real system) by grouping
many species. It represents the general pattern of carbon flow (an indicator
of food and energy) in the upper Chesapeake Bay during summer. Two basic
pathways dominate the estuarine food web. The direct pathway leads from
living plants to higher animals. The indirect, or detrital pathway leads from
dead organic matter to lower animals then to higher animals. Tidal marsh,
benthic, and submerged aquatic vegetation communities are strongly domi-
nated by the detrital pathway.
The following discussion outlines the components of the Chesapeake Bay
system and food web. Some of the primary producers of the Bay (plankton and
aquatic vegetation) and primary and secondary consumers (benthic organ-
isms, finfish, and waterfowl) are described in general terms.
PLANKTON
PHYTOPLANKTON AND BACTERIA
Phytoplankton are microscopic, usually single-celled plants, repre-
senting several divisions of algae. They constitute the base of the food chain;
the major primary producers in Chesapeake Bay. Thus, phytoplankton play a
fundamental role in the structure of the ecosystem. They are the major food
source for a number of species including zooplankton, benthic suspension
feeders, and fish. Bacteria are single-celled organisms that are responsible for
tremendous amounts of carbon and nutrient-cycling processes (see Figure 1).
As part of the detritus food chain, their role in decomposition of organic
matter, particularly dead plankton cells, is a major causative factor of anoxia
in bottom waters of the Bay.
In the surface waters of the Bay, dissolved nutrients and sunlight are
taken up by these photosynthetic organisms. Factors which control fluctu-
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ations in phytoplankton numbers, composition, and production are critical to
the success or failure of higher trophic levels. The balance among photo-
synthesis, nutrient exchange and predation ultimately determines planktonic
species composition. Large changes in nutrient and toxic loadings can also
cause changes in the quantity and quality (size and species composition) of
plankton communities in the system. There is growing evidence that a com-
bination of factors, probably arising from the synergistic effect of point and
nonpoint source discharges of toxics and nutrients, are causing a shift in
species composition. This shift is reflected in high production of bacteria and
minute phytoplankton species (favoring microzooplankton production) and
may be related to reduced population numbers in the higher trophic levels of
the system. Oysters, for example, may grow more slowly in areas where nu-
trient enrichment has shifted phytoplankton species composition to smaller
species which are not suitable as food.
ZOOPLANKTON
Zooplankton are swimming or floating animals that range from micro-
scopic to jellyfish size. Many are important food for fish and other organisms.
Zooplankton represent important primary consumers in the Chesapeake Bay
food web, and thus function as a key link in the transfer of energy derived
from phytoplankton, bacteria and detritius to higher trophic levels. Some
zooplankton, particularly the mesozooplankton (medium-size), function as
important and often critical links by supplying food to larval stages of many
fish and shellfish species in higher trophic levels. The distribution of meso-
zooplankton and the phytoplankton upon which they feed is a function of
salinity.
Jellyfish, including ctenophores (comb jellies) and sea nettles, prey on
the smaller zooplankton and may influence summer planktonic populations
and distributions. Microzooplankton, which are mostly single-celled protozoa,
feed heavily on bacteria. The larvae of benthic animals and fish are also
considered to be zooplankton. These larvae prey on smaller forms of plankton
and may be consumed by larger animals. As the larvae develop, they may in
turn consume other zooplankton.
VEGETATION
SUBMERGED AQUATIC VEGETATION
Submerged aquatic vegetation (SAV) is one of the Chesapeake Bay's most
significant natural resources. In 1976, the decline of SAV was selected as one
of the three major Bay problems (the only one directly focused on living
resources) to be further researched. Since that time, SAV has remained at the
forefront of public consciousness. It provides food and habitat for fish,
numerous other aquatic organisms, and waterfowl. SAV remains a visible
indicator of good water quality and the general ecological health of the
Chesapeake Bay.
Several of the key species identified for detailed analysis in this effort
require SAV (directly or indirectly) for food and/or habitat. Plants such as
eelgrass (a common SAV species in mid to high salinity regions) and emergent
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marsh grasses are major sources of primary productivity in the shallow waters
of the Bay. In addition to being a direct food source for some consumers,
organic detritus produced by decomposition of plant material provides food for
other primary consumers such as small crabs, shrimp, selected fish and other
detritivores.
Associations between SAV and finfish, shellfish, and waterfowl are well
documented. The most important waterfowl wintering areas have been the
most abundantly vegetated. Fish abundance in SAV communities in the upper
Bay is high, indicating the importance of SAV for food and shelter. Lower Bay
SAV beds serve as a primary blue crab nursery, sheltering large numbers of
juvenile blue crabs throughout the year.
Because prey organisms use SAV habitats, predators may be attracted to
the beds. Adult fish, such as striped bass and bluefish, may hunt invertebrate
prey in SAV beds. Summer resident wading and shore birds seek prey in or
near SAV beds.
SAV also functions as an important stabilizer for sediments. As turbid
water circulates through SAV beds, sediments tend to settle out, resulting in
clearer water and increased light transmittance. Direct uptake of nitrogen
and phosphorus by SAV and its associated epiphytes also serves to buffer
nutrient levels in the water during the spring and summer growing season.
Decomposition of SAV releases nutrients back to the water column during the
fall and winter when water column nutrient concentrations are lower.
TIDAL WETLANDS
The abundance of food and shelter provided by marsh grasses ensures a
very favorable habitat for other members of this community. A host of
invertebrates feed on decomposed plant material and, in turn, provide food for
numerous species of higher animals. Another source of food is the dense layer
of bacteria, algae, and microscopic animals that coats the stems of marsh
plants. Decomposing plants and, to a lesser extent, dead animals are major food
sources for the marsh dwellers. Therefore, the primary food web in the marsh
environment is based on detritus. Tidal marshes are also important as physical
habitat for estuarine species.
Salinity and frequency of tidal flooding are the most important factors in
determining the types of plant and animal populations that inhabit a par-
ticular marsh. Freshwater marsh vegetation includes cattails, reeds, arrow-
arum, big cordgrass, wild rice, three-square, tearthumb and pickerel weed.
Salt marshes of the mid and lower Bay are dominated by salt meadow cordgrass,
saltgrass, and saltmarsh cordgrass. Irregularly flooded salt marshes have the
fewest plant species and are dominated by needlerush.
Situated at the boundary between land and water, marshes absorb the
erosive energy of waves and may also act as nutrient buffers, regulating the
flow of local sources of nutrients into the Bay. Nutrients taken up by marsh
vegetation are later slowly released into the Bay during decomposition.
Marshes also protect the Bay ecosystem by trapping sediments that enter from
streams or tidal flooding.
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BENTHOS
The Chesapeake Bay supports an active community of organisms which
live in association with bottom sediments or attached to solid substrate such as
oyster shells, pilings, rocks, and shoreline structures. This assemblage,
collectively known as the benthos, represents a major component of the Bay
ecosystem. The benthos forms an important link between primary producers
and higher trophic levels. Many benthic organisms are principal food
sources for fish, waterfowl and crabs, while others are of direct economic
importance (crabs, clams, oysters). Benthic organisms also play a significant
role in the detrital pathway, breaking down organic matter. These decom-
posers are responsible for many key benthic processes, including nutrient re-
cycling, sediment chemistry, and the depletion of dissolved oxygen.
The temporal and spatial distribution of benthic communities is deter-
mined primarily by chemical and physical factors (mainly salinity, depth,
substrate, dissolved oxygen concentration, and temperature). The distribution
and abundance of organisms composing benthic communities are, therefore,
likely to respond to changes in water and sediment quality. Many benthic
organisms live for 1-2 years or longer so that benthic communities are
excellent indicators of an area's short and long-term trends in environmental
quality. In addition, because benthic organisms past the larval stage are rela-
tively immobile, they often complete much of their life cycles within well-
defined regions of the Bay. As a result, benthic responses to changes in
habitat quality are likely to be region-specific. As important intermediate
links in the Bay's food web, benthic community responses to habitat changes
are also likely to be representative of the responses of other living resources.
FINFISH
Finfish represent the majority of Chesapeake Bay nekton species. The
trophic relationships of fish are diverse, depending on developmental stage,
life histories, or physiological adaptations of different species. Most of the
large fish species of the Bay like bluefish, striped bass, and sea trout, are
temporary residents, living in the Bay for part of the year or only during
certain stages of their life cycles to spawn or feed. Resident finfish, such as
bay anchovies, hogchokers, and white perch, tend to be smaller in size. The
spawning behaviors of Chesapeake Bay finfish place them into two main cate-
gories: ocean-spawning fish (spot, croaker, menhaden) and freshwater or
estuarine-spawning fish (striped bass, herrings, shad).
Finfish occupy different trophic levels at specific stages of their lives.
Most finfish initially feed on zooplankton and later turn to larger prey. The
highest rates of survival of larval stages have been shown to correlate
positively with the highest zooplankton densities. Thus, the success of species
using the Bay as nursery grounds in its early life stages is dependent on the
availability of certain types of plankton.
Finfish are represented by all consumer levels within the Bay's food
web. Primary consumers, such as abundant schools of plankton-feeding
menhaden, represent a major pathway from the primary producers directly to
harvestable resources. Bluefish and striped bass are secondary or tertiary
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consumers, feeding on smaller finfish. Finfish also serve as prey for other
consumer-level species. The diets of many invertebrates, waterfowl, and some
mammals are composed largely of fish.
WATERFOWL AND WILDLIFE
In addition to the Chesapeake Bay's importance as a source of valuable
finfish and shellfish resources, the marshes and woodlands surrounding the
Bay provide habitat for a variety of waterfowl, birds and other vertebrates.
The Chesapeake Bay is part of an important migratory path known as the
Atlantic flyway. Most of the waterfowl reared between the western shore of
Hudson Bay and Greenland spend some time in the marshes and on the waters
of the Chesapeake Bay during their migrations. The Bay and the Delmarva
peninsula provide some of the prime, most heavily used waterfowl wintering
habitat along the Atlantic flyway.
Like finfish, bird species occupy all consumer levels of the food web.
Some birds feed on primary consumers (such as mollusks), while other species
feed on primary producers (plants). Birds feeding on secondary consumers,
such as fish, are considered tertiary consumers; at the extreme edge of the food
web, these high-level consumers (e.g. bald eagles) are often the first to be
affected by disruption of the ecological integrity of the Bay.
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CHESAPEAKE BAY HABITAT ZONATION
The variety of habitats within the Chesapeake can be classified using the
two most basic factors controlling the distribution of Bay biota: water depth
and salinity. In this classification of Bay habitats, gradients of depth and
salinity can be divided into descriptive zones. Depths range from the deepest
troughs and channels in the mainstem Bay to the intertidal shores and critical
land areas bordering tidal waters. Salinity ranges from the tidal freshwater
stretches of Bay tributaries and upper Chesapeake to the ocean-like water at
the mouth of the Bay. Within these zones, many other physical and biotic
factors such as sediment type, the presence of food and cover, the strength of
waves and currents, water temperature, dissolved oxygen, and habitat con-
tamination and disturbance control the distribution and abundance of living
resources. A generic system of habitat zones, defined in terms of salinity and
depth, offers a simplistic way to classify, describe, monitor, and manage living
resources in Chesapeake Bay.
Brief descriptions of depth and salinity zones follow, along with examples
of representative species in each -sone.
DEPTH ZONES
UPLAND SHORES
A variety of vegetation types exists on the upland shores which are the
terrestrial communities at elevations above the influence of tides. In many
cases, the physical nature of these upland regions is heavily influenced by
human activities, especially development and agriculture. Several species that
depend upon Bay aquatic habitats also rely upon these terrestrial environ-
ments for food, cover, or nesting sites. Examples of these species include the
bald eagle, Canada goose, river otter, beaver, and mink.
INTERTIDAL AND LITTORAL
The intertidal and littoral zones include areas with water depths of
approximately 0.5 meters (m) or less. They are semi-aquatic habitats, covered
periodically by tidal waters or washed by waves. These zones include marshes.
sandy beaches, mudflats, and shoreline structures such as revetments and
bulkheads. Representative species include marsh grasses, shorebirds, water-
fowl, muskrats, many benthic species, and larval or juvenile stages of finfish
and crabs.
SHALLOW WATER
The shallow water zone (to a depth of < 3 m) includes the uppermost
waters over the surface of the entire Bay and its tidal tributaries as well as the
bottom sediments in the shallow-water areas. Examples of important resident
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organisms include submerged aquatic vegetation, waterfowl, shallow-water
benthic species, crabs, and most juvenile finfish.
MID-WATER
The intermediate zone, with water depths between 3 and 6 m, includes the
mid-layer of pelagic waters and the underlying sediments. Submerged aquatic
vegetation is absent from all but the clearest waters at these depths. Oyster
bars and softshell clam habitat are most common in this zone. Oyster bars
support a specialized community of invertebrates, finfish and microorgan-
isms. In the summer, finfish, crabs, and other invertebrates which would
normally inhabit deeper water may be restricted to the intermediate zone by
the availability of dissolved oxygen.
DEEP WATER
Deep pelagic waters of the Bay having water depths of > 6 m constitute
habitat for most of the larger adult finfish. Many infaunal benthic species
inhabit the underlying sediments. Seasonal depletion of dissolved oxygen in
much of the Bay's deeper waters probably has limited the distribution of
species that otherwise would depend on these habitats. Examples include adult
striped bass, sciaenid finfish (croaker, spot, weakfish), flounder, sturgeon, and
infaunal invertebrates such as Macoma clam.
SALINITY ZONES
The absolute geographic location of salinity zones varies greatly, in-
fluenced by freshwater discharge, tides, weather, and water depth. Each
salinity zone includes the associated sediments and intertidal habitat.
TIDAL FRESH
The tidal fresh zone has salinities of < 0.5 ppt and includes the upper tidal
reaches of all Bay tributaries and the area of the upper Bay known as the
Susquehanna Flats. The tidal areas are critical spawning grounds for anadro-
mous finfish, but otherwise support mostly freshwater species of finfish,
invertebrates and plankton. Tidal fresh zone residents also include several
species of freshwater marsh plants, submerged aquatic vegetation, as well as
raptors, waterfowl, and upland wildlife.
OLIGOHALINE
The oligohaline zone, with a salinity range of 0.5 - 5.0 ppt, generally
includes the middle reaches of tidal tributaries and a portion of the upper
mainstem Bay, usually between the Susquehanna Flats and the mouth of the
Patapsco. These areas support fresh and brackish water species of aquatic
vegetation and are important nursery areas for anadromous finfish and
spawning grounds for estuarine finfish. Benthic species diversity is at its
lowest level in this zone, but some characteristic species (e.g. brackish-water
clam (Rangia cuneata)) are dependent upon it and can be present in high
densities. This zone is also characterized by high turbidity since it is a mixing
-12-
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zone of freshwater flow on the surface and the heavier, saline water along the
bottom.
MESOHALINE
The mesohaline portion comprises the most extensive salinity zone in the
Chesapeake Bay and has salinities ranging from 5.0 to 18 ppt. Under average
rainfall conditions, this zone encompasses the mainstem Bay from the mouth
of the Patapsco to the area just south of the Potomac River mouth. The lower
reaches of the major tributaries in the upper Bay are also mesohaline. Most of
the Chesapeake Bay species of finfish, shellfish and benthic organisms, along
with euryhaline (tolerant of a wide range of salinities) marine species,
inhabit this zone.
POLYHALINE
Most of the polyhaline zone, with salinity ranging from 18 to 32 ppt., is
found in the Virginia portion of the mainstem Bay. The lower reaches of the
York and James rivers are also in this zone. Some marine finfish live solely in
this segment of the Bay, although most of the estuarine finfish species are also
present. Spawning and overwintering habitat for female blue crabs occurs
within the polyhaline zone near the Bay mouth. Some benthic invertebrates
such as the hard clam (Mercenaria mercenaria), the whelk or "conch"
(Busycon spp.), and the oyster drill (Urosalpinx spp.), are generally restricted
to this zone. Saltmarsh grass (Spartina spp.), eelgrass (Zostera sp.), and
widgeongrass (Ruppia sp.) are typical in the polyhaline zone.
-13-
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SPECIES SELECTION
REPRESENTATIVE LIVING RESOURCES
The following list of species or species associations was developed by the
Living Resources Task Force to serve as an indicator of the Bay's ecological
condition. Not all species are indicators of recovery; rather, the abundance of
some are reflective of poor habitat conditions for less tolerant species. The list
includes species of commercial and recreational importance and species
which, due to their abundance, productivity, or distribution, are important in
the flow and accumulation of energy through various trophic levels of the
Chesapeake Bay ecosystem.
PHYTOPLANKTON ASSOCIATIONS:
Oligohaline
Winter/Spring
Cyclotella striata
Melosira granulata
Melosira islandica
Katodinium rotu.ndatu.rn
Cyclotella meneghiniana
Skeletonema costatum
Summer/Fall
Cyclotella striata
Merismopedia spp.
Microcystis aeruginosa
Gymnodinium spp.
Argetoceros spp.
Skeletonema costatum
Mesohaline
Winter/Spring
Skeletonema costatum
Cyclotella striata
Heterocapsa triquetra
Certaulina pelagica
Asterionella glacialis
Asterionella japonica
Summer/Fall
Cyclotella striata
Cryptomonas spp.
Skeletonema costatum
-15-
-------
Summer/Fall (continued)
Leptocylindrus minimus
Polyhaline
Winter/Spring
Skeletonema costatum
Leptocylindrus danicus
Asterionella glacialis
Cerataulina pelagica
Thalassiosira nordenskioldii
Thalassiosira rotula
Summer/Fall
Prorocentrum micans
Prorocentrum minimum
Heterocapsa triquetra
Cryptomonas spp.
Skeletonema costatum
ZOOPLANKTON ASSOCIATIONS:
Tidal fresh to oligohaline
Bosmina longirostris (Cladoceran)
Leptodora kindtii
Cyclops spp.
Mesocyelops edax
Diaptomus spp.
Tintinnids
Mesohaline to polyhaline
Winter
Cyanea capillata (lion's mane jellyfish)
Eurytemora affinis (copepod)
Acartia clausi (copepod)
Pseudocalanus spp.
Centropages hamatus
Temora longicornis
Neomysis americana
Sagitta elegans
Oithona spp.
Summer
Chrysaora quinquecirrha (sea nettle)
Mnemiopsis leidyi (ctenophore)
Podon polyphemoidese (cladoceran)
Evadne tergestina
Acartia tonsa (copepod)
Pseudodiaptomus coronatus
Labidocera aestiva
Parvocalanus crassirostris
Neomysis americana
-16-
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Summer (continued)
Sagitta tenius
Scottolana canadenis (meiobenthic copepod)
Ectinosonia centicorne (meiobenthic copepod)
SUBMERGED AQUATIC VEGETATION SPECIES:
Ruppia maritima (widgeongrass)
Zostera marina (eelgrass)
Vallisneria americana (wild celery)
Potamogeton pectinatus (sago pondweed)
Potamogeton perfoliatus (redhead grass)
EMERGENT AQUATIC VEGETATION SPECIES:
Spartina alterniflora (salt marsh cordgrass)
Spartina cynosuroides (big cordgrass)
Spartina patens (salt meadow cordgrass)
Juncus roemerianus
BENTfflC ASSOCIATIONS:
Tidal fresh
Tubificidae (Limnodrilidae)
Chironomidae
Corbicula manilensis (Asian clam)
Oligohaline
Rangia cuneata (brackish water clam)
Scolecolepides viridis (polychaete worm)
Mesohaline
Macoma balthica (Baltic clam)
Heteromastus filiformis (polychaete worm)
Streblospio benedicti (polychaete worm)
Leptocheirus plumulosus (amphipod)
Mya arenaria (soft-shelled clam)
Polyhaline
Loimia medusa
Mulinia lateralis
Asabellides oculata
Sphiophanes bombyx
Mercenaria mercenaria (hard clam)
Maldanids
Tellinids
Nephtyiids
Phoxocephalids
Haustoriids
-17-
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Euryhaline
Callinectes sapidus (blue crab)
Motile epifauna
Palaemonetes pugio (grass shrimp)
Gammarus gammarus (amphipod)
Crangon
Corophium
Mysidacea
Sessile epifauna
Balanus improvisus (barnacle)
Mytilis edulis
Molgula spp.
Bryozoa
Crassostrea virginica (American oyster)
Anemones
FINFISH SPECIES:
Freshwater and Estuarine Spawners
Alosa sapidissima (American shad)
Alosa pseudoharengus (alewife)
Alosa aestivalis (blueback herring)
Alosa mediocris (hickory shad)
Anchoa mitchilli (Bay anchovy)
Menidia menidia (Atlantic silverside)
Morone saxatilis (striped bass)
Morone americana (white perch)
Perca flavescens (yellow perch)
Acipenser oxyrynchus (Atlantic sturgeon)
Acipenser brevirostrum (shortnose sturgeon)
Fundulus heteroclitus (mummichog)
Micropterus salmoides (largemouth bass)
Pseudopleuronectes americanus (winter flounder)
Trinectes maculatus (hogchoker)
Cynoscion regalis (weakfish)
Cynoscion nebulosus (spotted seatrout)
Pogonias cromis (black drum)
Ocean Spawners
Brevoortia tyrannus (menhaden)
Leiostomus xanthurus (spot)
Micropogonias undulatus (Atlantic croaker)
Sciaenops ocellatus (red drum)
Centropristis striata (black sea bass)
Paralichthys dentatus (summer flounder)
Pomatomus saltatrix (bluefish)
Anguilla rostrata (eel)
-18-
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WATERFOWL AND OTHER AQUATIC BIRD SPECIES:
Anas platyrhynchos (mallard)
Anas rubripes (black duck)
Aythya valisneria (canvasback)
Aythya americana (redhead duck)
Aix sponsa (wood duck)
Ardea herodias (great blue heron)
Florida caerulea (little blue heron)
Butorides striatus (green-backed heron)
Casmerodius albus (American egret)
Egretta thula (snowy egret)
Pandion haliaetus (osprey)
Haliaeetus leucocephalus (bald eagle)
Clangula heimalis (old squaw)
Melanitta deglandi (white-winged scoter)
Olor columbianus (tundra swan)
Megaceryle alcyon (kingfisher)
Anas acuta (northern pintail)
Anas strepera (gadwall)
Anas americana (American widgeon)
Branta canadensis (Canada goose)
Sterna albifrons (least tern)
Haematopus palliatus (oystercatcher)
Rynchops niger (black skimmer)
Limnodromus spp. (dowitcher)
Arenaria interpres (ruddy turnstone)
Actitis macularia (spotted sandpiper)
OTHER VERTEBRATE SPECIES:
Mustela vison (mink)
Lutra canadensis (river otter)
Ondatra zibethica (muskrat)
Castorcanadensis (beaver)
Caretta caretta (Atlantic loggerhead turtle)
Lepidochelys kempi (Atlantic ridley turtle)
Malaclemys terrapin (diamondback terrapin)
TARGET SPECIES
The following list of target species, selected from the list of key repre-
sentative species by the Living Resources Task Force, was reviewed by partici-
pants at the Habitat Requirements Workshop held on February 24, 1987. Selec-
tion criteria are outlined in the introduction of this document. Species
grouped together with the symbol "*" were determined to have habitat
requirements similar enough to permit treatment as a group rather than as
individuals.
-19-
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SUBMERGED AQUATIC VEGETATION:
FINFISH:
Ruppia maritima (widgeongrass)
Zostera marina (eelgrass)
Vallisneria americana (wild celery)
Potamogeton pectinatus (sago pondweed)
Potamogeton perfoliatus (redhead grass)
Morone saxatilis (striped bass)
* Alosa aestivalis (blueback herring)
* Alosa pseudoharengus (alewife)
* Alosa sapidissima (American shad)
* Alosa mediocris (hickory shad)
Perca flavescens (yellow perch)
Morone americana (white perch)
Brevoortia tyrannus (menhaden)
Leiostomus xanthurus (spot)
Anchoa mitchilli (bay anchovy)
SHELLFISH:
Molluscan
* Crassostrea virginica (American oyster)
* Mya arenaria (softshell clam)
* Mercenaria mercenaria (hard clam)
Crustacean
Callinectes sapidus (blue crab)
WATERFOWL AND OTHER AQUATIC BIRDS:
Aythya americana (redhead duck)
Anas rubripes (black duck)
Aythya valisneria (canvasback)
Aix sponsa (wood duck)
* Ardea herodias (great blue heron)
* Florida caerulea (little blue heron)
* Butorides striatus (green-backed heron)
* Casmerodius albus (American (great) egret)
* Egretta thula (snowy egret)
* Pandion haliaetus (osprey)
* Haliaeetus leucocephalus (bald eagle)
-20-
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HABITAT MATRICES
The Living Resources Task Force, aware of the many limitations and gaps
in the available information, has summarized minimum habitat requirements
for selected target species. The abundance and diversity of the Bay's living
resources are affected by several variables, many of which are not fully
understood. If the recovery of species which have declined in the Chesapeake
Bay and the reestablishment of a more balanced ecosystem are the ultimate
measures of success, the achievement of certain minimum habitat require-
ments for specific regions in the Chesapeake Bay is an essential first step.
The following text and matrices summarize existing information on habi-
tat requirements for the initial list of target species. For many species, reli-
able in situ water quality and habitat requirements are not known and numer-
ous data gaps exist. In all instances, the Living Resources Task Force reviewed
available laboratory and field studies which evaluated the tolerance of species
to individual variables such as salinity, turbidity, dissolved oxygen, and toxics.
Few studies dealt with the composite effects of water quality and habitat factors
on survival. These variables are closely interrelated and a change in one
variable often affects the relative tolerance to other factors. Water temper-
ature, for example, is inversely proportional to dissolved oxygen. Since rates
of respiration rise with increasing water temperature, animals can tolerate
lower oxygen concentrations longer at lower temperatures. Toxic substances
demonstrate similar interactions. In combination, these materials can exert
either synergistic or antagonistic effects and their relative toxicity is gen-
erally inversely proportional to dissolved oxygen. When such interactions
could clearly be identified, they have been noted in the text or accompanying
matrices.
The matrices contain information available for the sensitivities of target
species to toxic substances. The sensitivities have been included in the form in
which they were reported in the literature (LC50, LCD, etc.). These should not
be construed as levels of toxic materials that will necessarily protect the
resources. Future efforts must address the interpretation of existing toxics
data in the determination of specific habitat requirements.
The following sections describe the necessary requirements for each
target species.
TARGET SPECIES GROUP: Submerged aquatic vegetation complex
Critical life stage: all life stages
Critical period: April-September
Five species of submerged aquatic vegetation (SAV), with tolerances
spanning the full range of salinities found in Chesapeake Bay habitats, were
-21-
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selected as members of the target species group. Widgeongrass (R up pi a
maritima) and eelgrass (Zostera marina) are representative of both the meso-
haline and polyhaline zones. Sago pondweed (Potamogeton pectinatus) and
redhead grass (P. perfoliatus) are tolerant of oligohaline and mesohaline
salinities. Wild celery (Vallisneria americand) inhabits tidal fresh and oligo-
haline waters.
Submerged aquatic plants are particularly appropriate as target species
because of their key role in providing critical habitat for other species. An
SAV bed provides cover for fish and invertebrates, food for waterfowl and
reduces shore erosion and suspended sediment loads. Also, SAV is a good
indicator of poor water quality due to its sensitivity to turbidity and nutrient
enrichment.
Light penetration limits the depth at which SAV can survive and grow.
In Chesapeake Bay, this depth is usually less than 2 m, although in less turbid
water some SAV species may grow at depths of 6 m or more. Dense
phytoplankton blooms and epiphytic growth, stimulated by high nutrient
levels, can reduce the transmittance of light to SAV leaves. Shading reduces
photosynthetic activity causing depletion of carbohydrate reserves required
for growth, reproduction, and overwintering. In high salinity waters, ni-
trogen is generally a limiting nutrient. High nitrogen concentrations can
cause phytoplankton blooms and epiphytic growth harmful to SAV. In the
mesohaline zone, either nitrogen or phosphorus can limit algal growth.
Levels of dissolved inorganic nitrogen greater than 0.14 mg/1 and dissolved
inorganic phosphorus greater than 0.01 mg/1 are thought to be responsible
for previous SAV declines, largely because of excessive epiphytic growth and
high algal concentrations in surrounding waters (Stevenson, unpublished
data).
Suspended sediment also can limit light penetration in the water column.
Light attenuation coefficients (kd) for photosynthetically active radiation
(400-700 nm wavelength) should not exceed 2.0/m, and total suspended solids
should be less than 20 mg/1 to promote reestablishment of SAV (Figure 2)
(Stevenson, unpublished data) in mesohaline zones.
Substantial regrowth of SAV in the tidal fresh portion of the Potomac
River has been attributed to recent reductions in phosphorus loadings from
the Blue Plains sewage treatment plant. In freshwater at the head of the Bay,
SAV grows well in the presence of high nitrate levels apparently because
phosphate concentrations are low enough to limit phytoplankton growth. In
these areas, SAV is able to obtain sufficient phosphorus from the sediments.
Dense beds of some SAV species, however, can raise daytime pH levels high
enough to cause chemical reactions which act to release phosphate from sedi-
ments, stimulating algal growth.
Herbicides, such as atrazine, can be harmful to SAV at concentrations in
excess of 10 ug/L. Water column concentrations of this magnitude are likely to
occur in localized shallow embayments directly affected by agricultural
runoff.
-22-
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TARGET SPECIES: Striped bass (Morone saxatilis)
Critical life stage(s): larval, juvenile
Critical life period: April to June
BACKGROUND
There have been numerous literature reviews and synopses dealing with
striped bass biology (e.g. Richkus, 1986; Setzler-Hamilton, 1980; Westin and
Rogers, 1978; and Hildebrand and Schroeder, 1928). The reader is referred to
these publications for a more thorough account of their life history.
SPAWNING AND RANGE
Striped bass spawn during the spring in tidal fresh or brackish waters.
The principal spawning and nursery areas of striped bass along the Atlantic
Coast are found in the Chesapeake Bay and its tributaries (Merriman, 1941) and
the Hudson and Roanoke rivers (Kaumeyer and Setzler-Hamilton, 1982).
Within the Chesapeake Bay basin, major spawning areas include: the
James, Pamunkey, Mattaponi. Rappahannock, Patuxent, and Potomac rivers on
the western shore; the head of the Bay with the Susquehanna Flats, Elk River,
Chesapeake and Delaware (C & D) Canal; and, the Choptank and Nanticoke
rivers on the Eastern Shore (Mansueti and Hollis, 1963; Speir, Personal
Communication, 1987).
Spawning activity is apparently triggered by a rise in water temperature.
Spawning times may vary from year to year due to annual temperature vari-
ations. In the Chesapeake Bay, 1 to 3 peaks occur during each spawning
season with the major peak occurring any time during the last half of April or
the first week of May (Kaumeyer and Setzler-Hamilton, 1982; Grant and Olney,
1982). Research has suggested that freshwater flow (both velocity and
volume) is related to successful spawning (Kaumeyer and Setzler-Hamilton,
1982; Bayliss, 1982).
TROPHIC IMPORTANCE
Adult and copepodite copepods and cladocerans are the major food items of
larval striped bass. Setzler-Hamilton et al. (1981) reported that rotifers and
Eurytemora affinis copepodites are the dominant prey for first-feeding striped
bass larvae in the Potomac River. Larval striped bass from 6 to 13 mm consume
copepodites, adults of cyclopoids and other copepods. The diet of larvae >. 14 mm
consists almost entirely of adult copepods (Kaumeyer and Setzler-Hamilton,
1982). Westin and Rogers (1978) provided a comprehensive list of food items
for striped bass at various life stages.
TOXICITY
Of all the species examined in this report, striped bass has been studied
the most with respect to its sensitivity to toxic chemicals. This section sum-
marizes selected striped bass bioassays and highlights conflicting data.
-24-
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Hall (1984) reported that water quality data from an on-site toxicity
experiment on the Nanticoke River implicated that aluminum toxicity was
induced by low pH. According to Richkus (1986), striped bass exhibited "no
detectable effect" from aluminum concentrations of 200 to 400 ug/1 at about pH
7. However, a pH of 6.5 or less with aluminum concentrations in the range of
25 to 100 ug/1 caused significant mortality dependent upon the life stage of the
striped bass (Richkus, 1986). O'Rear (1972) compared the relative toxicity of
copper and zinc on embryos. Copper was more toxic, with a 48 hr LC50 value of
0.74 ppm. Hughes (1973) tested the tolerance of larval striped bass to cadmium,
copper, and zinc. Cadmium was the most toxic. Larval striped bass experienced
50% mortality when exposed to 0.001 ppm of cadmium chloride for 96 hr
(Kaumeyer and Setzler-Hamilton, 1982).
Data indicate that levels of total residual chlorine (TRC), while not neces-
sarily lethal, may have significant sublethal effects on striped bass. For
example, striped bass larvae exhibited significantly shorter body lengths after
eggs were exposed to 0.15 ppm of total residual chorine. Kaumeyer and Setzler-
Hamilton (1982) report that striped bass eggs exhibit 50% and 100% reduction
in hatch rate when exposed to 0.19 and 0.43 ppm of TRC, respectively.
Lethal concentrations of toxic substances at various stages of the striped
bass life history have been summarized by Richkus, 1986; Westin and Rogers,
1978; DiNardo et al., 1984; Emergency Striped Bass Study, 1984; and, Bonn et al.,
1976.
Appendix A contains additional information on the sensitivity of striped
bass for a selected group of toxic substances.
TARGET SPECIES: Alewife (Alosa pseudoharengus)
Critical Life Stage(s): egg, larval
Critical Life Period: Early April to mid-June
TARGET SPECIES: Blueback herring (Alosa aestivalis)
Critical Life Stage(s): egg, larval
Critical Life Period: Early April to end of May
BACKGROUND
This profile covers the life history and environmental requirements of
the blueback herring (Alosa aestivalis) and the alewife (Alosa pseudo-
harengus), since their distributions overlap and their morphology, ecological
roles, and environmental requirements are similar. The alewife and blueback
herring are anadromous species found in riverine, estuarine, and Atlantic
coastal habitats, and have occurred historically throughout the Chesapeake
Bay region (Hildebrand and Schroeder, 1928). Since the early developmental
stages of the blueback herring, alewife, and hickory shad (Alosa mediocris)
are difficult to separate and the spawning seasons and locations overlap for all
these species, the matrix developed for both species also is applicable to the
hickory shad.
-26-
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SPAWNING AND RANGE
The spawning locations and seasons of blueback herring and alewife
overlap considerably. Blueback herring usually do not ascend streams as far
as alewives (Hildebrand and Schroeder, 1928; Scott and Grossman, 1973). Blue-
back spawn in both fresh and brackish water in rivers and ponds (Davis, 1973;
Hildebrand, 1963). However, Loesch and Lund (1977) reported that blueback
herring preferred spawning in fast-flowing waters with hard substrates.
Alewife often spawn in slower-moving waters (Wang and Kernehan, 1979).
Because spawning by blueback herring is more site-specific than for alewife,
dams and alteration of blueback spawning sites may be more detrimental to
their population.
The spawning period for these two species is also very similar. Blueback
spawning occurs from late April to early May in the Potomac River
(Hildebrand, 1963). Alewives spawn from early April through mid-May (Wang
and Kernehan, 1979).
Smith (1971) observed blueback spawning at water temperatures of 19-24
degrees C, but Wang and Kernehan (1979) reported slightly lower spawning
temperatures (15.0-22.0 degrees C). Alewives spawn at water temperatures
from 12.0-22.5 degrees C (Wang and Kernehan, 1979). Alewife eggs hatch at
temperatures ranging from 12.7-26.7 degrees C (Atlantic States Marine
Fisheries Commission, 1985). Klein and O'Dell (1987) report that the optimum
temperature range for river herring larvae is 16-24 degrees C.
TROPHIC IMPORTANCE
The river herrings, blueback herring and alewife, are seasonally abun-
dant fish feeding chiefly on zooplankton, particularly copepods (U.S. Corps of
Engineers, 1984). The larvae for these two species consume primarily zoo-
plankton and relatively small cladocereans and copepods (U.S. Fish and
Wildlife Service, 1983). Juveniles and adults consume fish, crustacean and
insect eggs, as well as adult insects; young fish may also constitute a portion of
the diet when available (U.S. Corp of Engineers, 1984).
ENVIRONMENTAL CONDITIONS
The LC50 of total residual chlorine (TRC) for blueback herring eggs
ranges from 0.20-0.32 ppm (U.S. Fish and Wildlife Service, 1983). Eggs exposed
to 84 mg/1 of TRC reached early embryo stages but failed to develop further.
Larvae from eggs exposed to sublethal concentrations of total residual
chlorine were all deformed. Concentrations of 36 mg/1 TRC produced 100%
mortality in 1-day old larvae (U.S. Fish and Wildlife Service, 1983). Ammonia,
nitrites and any form of reduced nitrogen are toxic. Nitrogen and phosphorus
can indirectly affect food production and induce anoxic conditions (Connery,
1987).
Auld and Schubel (1978) found that suspended sediments at concentra-
tions of 100 ppm or less had no significant effect on the hatch rate of alewife
or blueback herring eggs. Research suggests that water flow created by
shear, power plant uptake, pressure drop, and dam turbines is critical to the
reproduction and survival of river herrings (Connery, 1987).
-27-
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TARGET SPECIES: American shad (Alosa sapidissima )
Critical Life Stage(s): egg, larval
Critical Life Period: Mid-April to early June
TARGET SPECIES: Hickory shad (Alosa mediocris )
Critical Life Life Stage(s): egg, larval
Critical Life Period: April to June
BACKGROUND
Historically, shad have inhabited virtually all rivers feeding the
Chesapeake Bay (Kaumeyer and Setzler-Hamilton, 1982). Currently, shad pop-
ulation numbers are extremely low in Maryland waters, and shad fishing is
banned (Jones et al., 1978; Kaumeyer and Setzler-Hamilton, 1982). There is
still a commercial shad fishery in Virginia tributaries, however.
SPAWNING AND RANGE
Spawning runs may begin as early as February, but are most frequent in
April. Characteristic spawning and nursery grounds for shad are tidal fresh-
waters in estuaries and rivers; however, some shad can tolerate moderate
salinities (Stagg, 1985; Kaumeyer and Setzler-Hamilton, 1982). Successful
hatches have been reported at salinities ranging from 7.5 ppt at 12.0 degrees C
to 15 ppt at 17 degrees C. No eggs hatched at a salinity of 22.5 ppt (U.S. Fish and
Wildlife Service, 1986).
Shad spawning areas vary in depth and substrate. Shad seem to prefer
areas dominated by shallow water or broad flats with sand or gravel bottoms
(U.S. Fish and Wildlife Service, 1986). Sufficient water current velocities are
required to keep the shad eggs suspended in the water column. Preferred
velocities in spawning waters range from 30.5 to 91.4 cm/sec (U.S. Fish and
Wildlife Service, 1986). Exposure of the eggs to suspended sediment concen-
trations as high as 1,000 mg/1 did not affect hatching success (Auld and
Schubel, 1978), but larval mortality was high at suspended sediment concen-
trations greater than 100 mg/1 for 96 hours (U.S. Fish and Wildlife Service,
1986).
ENVIRONMENTAL CONDITIONS
Eggs hatch in 12 to 45 days at 12 degrees C and in 6 to 8 days at 17 degrees C
(Bigelow and Schroeder, 1953). Maximum survival of eggs and larvae occurs at
15.5-26.6 degrees C (U.S. Fish and Wildlife Service, 1986). Temperatures of 7-9
degrees C were reported to be lethal to eggs and larvae and temperatures of
20.0-23.4 degrees C caused extensive larval abnormalities (U.S. Fish and
Wildlife Service, 1986). The LD50 for acid pH was 5.5 and it was 9.5 for basic pH
(U.S. Fish and Wildlife Service, 1986). Larval shad LD50 for low dissolved oxy-
gen (DO) ranges from 2.0-3.5 ppm, depending on the population. Mortality of
eggs was 100% at DO levels below 1.0 mg/1 (U.S. Fish and Wildlife Service, 1986).
Larvae exhibit significant signs of stress when exposed to a DO level of 3.0
mg/1, and many died at 2.0 mg/1 (Chittenden, 1969). A DO level of > 5.0 ppm is
considered optimum (Chittenden, 1969; Wang and Kernehan, 1979).
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Larvae remain near the spawning grounds, usually a short distance
downstream. Young remain in the nursery area until water temperatures
begin to decrease in the fall. The downstream migration begins at a water
temperature of approximately 21.1 degrees C (Wang and Kernehan, 1979). All
young have left the nursery grounds by the time the temperature reaches 8.3
degrees C (Wang and Kernehan, 1979).
TROPHIC IMPORTANCE
Shad larvae consume cyclopoid copepods, midge larvae, midge pupae, and
Daphnia pulex (U.S. Fish and Wildlife, 1986).
ADDITIONAL INFORMATION
For a concise overview see Boreman (1981); for a detailed study of the life
history of shad see Mansueti and Kolb (1953). Reports by Cooper (1984),
Richkus and DiNardo (1984), and Davis (1973) respectively provide thorough
reviews on the status of Atlantic coast shad, all anadromous alosids of the
eastern United States, and shad life history information for Virginia waters.
TARGET SPECIES: Yellow perch (Perca flavescens)
Critical life stage: egg, larval
Critical life period: first year of life
SPAWNING AND RANGE
Yellow perch make vertical temperature-dependent migrations and in-
shore, upstream spawning migrations. The spawning period lasts from March
to April in shallow tidal and non-tidal freshwater. Spawning occurs in low
velocity currents (< 5 cm/s). The species is common where debris or vege-
tation are present. Eggs are gelatinous and semibuoyant (U.S. Corps of
Engineers, 1984; U.S. Fish and Wildlife Service, 1983; and, Wang and Kernehan,
1979). In the Chesapeake Bay, yellow perch habitat is situated between the up-
stream limit of tidal freshwater to mid-mesohaline salinity zones. Spawning
activity has been reported in low salinity waters up to 2.5 ppt in the Severn
River (Wang and Kernehan, 1979). Hildebrand and Schroeder (1982) observed
yellow perch from Havre de Grace, Maryland to Lewisetta, Virginia. The fish
tend to migrate toward the shorezone in summer and into deeper waters in
winter (U.S. Corps of Engineers, 1984).
TROPHIC IMPORTANCE
The principal foods of young yellow perch in freshwater consists of
insects and small crustaceans (U.S. Corps of Engineers, 1984). Adults feed on
soft-bodied fish, minnows, and anchovies, as well as isopods, amphipods,
shrimp, crabs, insect larvae, and snails (U.S. Corps of Engineers, 1984; Hilde-
brand and Schroeder, 1928).
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OTHER SENSITIVITIES
Yellow perch inhabit slow-flowing tidal rivers containing vegetation,
submerged trees or pilings. Data suggest that yellow perch abundance de-
creases with increasing turbidity (U.S. Fish and Wildlife Service, 1983). They
are able to tolerate low dissolved oxygen levels and remain active even under
winter ice. However, laboratory and field studies determined that dissolved
oxygen levels from 0.2-1.5 mg/1 are lethal to yellow perch. A dissolved oxygen
level of 5 mg/1 was determined as the optimum lower limit (U.S. Fish and
Wildlife Service, 1983).
TARGET SPECIES: White perch (Morone americand)
Critical life stage(s): egg, larval
Critical life period: first year of life
BACKGROUND
White perch are found throughout the Chesapeake Bay and C&D Canal
and have been reported in marine waters north of Chesapeake Bay. White
perch are considered anadromous, but non-migratory resident populations do
occur.
SPAWNING AND RANGE
White perch move upriver in the spring into the shorezone of tidal fresh
waters to spawn (U.S. Corps of Engineers, 1984). In the Chesapeake Bay,
spawning occurs from April to June. Spawning has been observed in Decem-
ber when appropriate climatic conditions occurred (Hildebrand and
Schroeder, 1928). The species prefers spawning over shoal hard bottoms (e.g.
sand or gravel) with currents. During their first year, juveniles remain in
soft-bottomed, shallow, freshwater nursery areas, preferably in vegetated
zones. Juveniles larger than 25 mm in total length begin inshore-offshore
movements in response to light levels. Low temperatures cause white perch to
move into deeper waters. Wintering populations are found in the deeper
channels and holes in the upper Bay and tributaries. White perch in the Bay
system are thought to consist of isolated subpopulations indigenous to each
tributary.
Adult white perch are found in salinity zones of 5-18 ppt; however, they
prefer to spawn at salinities less than 4.2 ppt (U.S. Fish and Wildlife Service,
1983; U.S. Corps of Engineers, 1984). Osmotic regulation is disrupted in eggs de-
posited in water of salinities >_ 10 ppt. Larvae can tolerate salinities in the
range of 0-8 ppt (U.S. Fish and Wildlife Service, 1983).
TROPHIC IMPORTANCE
The white perch is a generalized feeder and is benthophagus or pisciv-
orous depending upon food availability, age and season (U.S. Fish and Wildlife
Service, 1983). Larvae prey upon zooplankton. Fish, crustaceans, annelids
and insect larvae are taken during juvenile and adult stages (Hildebrand and
Schroeder, 1928). The fry are consumed by larger prey fish such as bluefish
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and striped bass (Hildebrand and Schroeder, 1928; U.S. Fish and Wildlife
Service, 1983; U.S. Corps of Engineers, 1984).
TARGET SPECIES: Menhaden (Brevoortia tyrannus)
Critical life stage(s): juvenile
Critical life period: April to October
SPAWNING AND RANGE
Juvenile menhaden are found in upper Chesapeake Bay tributaries from
late May through November. Kaumeyer and Setzler-Hamilton (1982) report
that juveniles were found in the Potomac River in March and April and in the
upper Bay from late May through late June and in November. April through
October is generally the peak time of abundance in the upper Chesapeake Bay.
During the post-larval stage, menhaden tend to accumulate at the fresh/salt-
water interface in the upper Bay region. Juveniles in the upper Bay begin to
emigrate, generally after their first summer, from the freshwater interface
into the mesohaline zone (U.S. Corps of Engineers, 1984; Kaumeyer and Setzler-
Hamilton, 1982). Larger fish are found in the deeper waters down the Bay.
Sub-adults leave the estuary with the adults in October; however, some over-
wintering occurs in Chesapeake Bay (U.S. Corps of Engineers, 1984; Kaumeyer
and Setzler-Hamilton, 1982).
Spawning and early larval development occur in continental shelf waters
of the Atlantic. Menhaden are estuarine dependent, utilizing the estuary both
as a nursery for juveniles and as adult feeding ground during the summer
months (Bigelow and Schroeder, 1953; Reintjes, 1969; and U.S. Corps of
Engineers, 1984). Reintjes (1969) observed eggs and small larvae in Long
Island Sound, Narragansett Bay, and Chesapeake Bay, but suggested that
spawning in these areas made minor contributions to total population
numbers.
TROPHIC IMPORTANCE
Menhaden represent a major energy link between plankton directly to
the large piscivores. Where menhaden are present in dense schools, their
filter-feeding can be a primary control over local plankton abundance. Ac-
cording to Ulanowicz and Baird (1986), the summer diet of menhaden in the
mesohaline part of Chesapeake Bay consists of zooplankton (65%), phyto-
plankton (5%), and unspecified organic particulates (29%).
TARGET SPECIES: Spot (Leiostomus xanthurus)
Critical life stage(s): juvenile
Critical life period: Early April to early November
SPAWNING AND RANGE
The spot is a demersal, marine spawning fish. Spawning activity on the
continental shelf adjacent to the Chesapeake Bay was reported to occur during
late fall and winter (Kaumeyer and Setzler-Hamilton, 1982). Some adults may
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spawn twice a year (U.S. Fish and Wildlife Service, 1982). Kaumeyer and
Setzler-Hamilton (1982) suggested that adult spot do not survive after they
spawn.
Post-larval and juvenile spot spend much of their lives in estuaries (U.S.
Fish and Wildlife Service, 1982). Post-larval spot inhabit Chesapeake Bay from
early April through early November (Hildebrand and Schroeder, 1928). In the
Maryland portion of the Bay, spot larvae and young juveniles congregate in
the oligohaline zone, although when population densities are high, some
young move into tidal freshwater, shallow marshes, and drainage ditches (U.S.
Corps of Engineers, 1984; U.S. Fish and Wildlife Service, 1982). In the lower
Bay, spot larvae and young juveniles are found in mesohaline and polyhaline
tidal marshes. Spot are common near grass beds and over muddy substrates
(U.S. Fish and Wildlife Service, 1982). In Chesapeake Bay, adults are found in
mesohaline to polyhaline salinity zones (U.S. Corps of Engineers, 1984; U.S.
Fish and Wildlife Service, 1982). Spot leave the Bay as water temperatures de-
cline in the fall (Wang and Kernehan, 1979).
Fish in their second or third year of life do not penetrate very far into
the estuary, and are abundant only in the lower Virginia portion of the Bay
(U.S. Corps of Engineers, 1984). Adult spot habitat in the Chesapeake is defined
as mid-mesohaline to polyhaline areas with depths to 6 m overlying soft sedi-
ment bottoms (U.S. Corps of Engineers, 1984).
TROPHIC IMPORTANCE
Juvenile spot primarily consume benthic invertebrates including: ostra-
cods, copepods, and polychaetes (U.S. Fish and Wildlife Service, 1982).
Approximately 93% of the summer diet consists of polychaetes; most of the
remainder is Macoma spp. (Ulanowicz and Baird, 1986). Spot are preyed upon
by large gamefish and also harvested by sport and commercial fisheries. Spot
represent a significant link in the transfer of energy from the detritivores
and primary consumers eaten by spot in the Bay to its predators in the waters
of the adjacent continental shelf (U.S. Corps of Engineers, 1984).
TARGET SPECIES: Bay anchovy (Anchoa mitchilli)
Critical life stage(s): larval
Critical life period: May to September
BACKGROUND
Bay anchovy has been observed in virtually all open waters throughout
the Chesapeake Bay from the tidal fresh to the polyhaline zone; the C & D Canal
and Havre de Grace down to Lynnhaven Roads, Virginia (Wang and Kernehan,
1979; Hildebrand and Schroeder, 1928). Anchovy larvae are pelagic and are
also found over a wide salinity range (Wang and Kernehan, 1979; Hildebrand
and Schroeder, 1928). According to Wang and Kernehan, (1979) the larvae
move upstream to low salinity regions after hatching, with the highest con-
centrations of larvae observed at salinities of 0-7 ppt salinity. The U.S. Corps of
Engineers (1984) reported larvae at salinities of 3-7 ppt. Larvae were found 40
miles above brackish water in Virginia (Wang and Kernehan, 1979) and in the
Potomac River in freshwater near Bryans Point, about 12 miles below Wash-
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ington, D.C. (Hildebrand and Schroeder, 1928). Anchovy larvae also occur in
large numbers throughout the lower Chesapeake Bay (Olney, 1983).
SPAWNING AND RANGE
The Bay anchovy spawning season occurs from May to September in the
Chesapeake Bay (Wang and Kernehan, 1979). Spawning is pelagic and occurs
in the Chesapeake Bay at salinities ranging from 1-22 ppt (U.S. Corps of
Engineers, 1984; Wang and Kernehan, 1979). Spawning also occurs at the
Chesapeake Bay mouth where salinities are typically 25-28 ppt (Olney, 1983).
Wang and Kernehan (1979) reported that spawning activity in the Delaware
Bay occurs between 15 degrees C and 30 degrees C with peak activity occurring
at 22-27 degrees C. They also reported peak egg densities occur at salinities of
12-13 ppt in Chesapeake Bay. In the upper Chesapeake Bay, larvae are
observed in shallow shore areas where the salinities range between 3-7 ppt
(U.S. Corps of Engineers, 1979).
TROPHIC IMPORTANCE
Anchovies feed primarily on mysids and copepods (Hildebrand and
Schroeder, 1928). In overlapping ranges, Bay anchovy larvae are reported to
compete with alosid larvae for copepods (U.S. Corps of Engineers, 1984; Hilde-
brand and Schroeder, 1928). The anchovy is a year-round resident, and an
important forage fish of the Chesapeake (U.S. Corps of Engineers, 1984).
During the summer, in the mesohaline portion of Chesapeake Bay, anchovies
consume large quantities of phytoplankton (13%), zooplankton (72%), and
organic detritus (15%) (Ulanowicz and Baird, 1986).
ADDITIONAL INFORMATION
The larval stage is considered the most sensitive life stage for the Bay
anchovy. The larvae have been observed to congregate at the surface waters
of the oligohaline zone. Crowding has been observed as anchovies move into
the narrower oligohaline areas of tributaries. Concentration of larvae in the
surface waters may cause localized overpopulation which possibly resulting in
a reduction in year class abundance (U.S. Corps of Engineers, 1984).
TARGET SPECIES GROUP: Molluscan Shellfish
American oyster (Crassostrea virginica)
Critical life stage(s): larval, spat and adult
Critical life period: entire life cycle
Soft clam (Mya arenaria)
Critical life stage(s): larval
Critical life period: May - October
Hard clam (Mercenaria mercenaria )
Critical life stage(s): egg and larval
Critical life period: first year of life
-43-
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BACKGROUND
American oysters, soft clams, and hard clams are prominent members of
the benthic community in Chesapeake Bay and contribute substantially to the
economy of the region. Oysters have recently experienced severe declines in
abundance. Soft clams in the Chesapeake Bay have also decreased in abun-
dance in recent years in the Bay. Intense fishing pressure, loss of habitat, and
water quality degradation have been blamed for declines in the abundance of
these species. Hard clams, however, have maintained more stability in popu-
lation numbers, primarily due to greater market demand for surf clams and
ocean quahogs in the mid-Atlantic region.
SPAWNING AND RANGE
All Chesapeake oysters are subtidal, whereas their southern counterparts
are often intertidal. American oysters prefer a firm substrate: pilings, hard
rock bottoms, and substrates firmed with the oyster shells of previous gener-
ations. Soft clams in the Chesapeake inhabit shallow subtidal (10 m) estuarine
waters to intertidal areas in the oligohaline through the polyhaline zones.
Hard clams are euryhaline marine species sensitive to salinities below 12 ppt,
and thus are only found in the lower Bay from the mesohaline through the
polyhaline zone (12-32 ppt). Although found in a variety of substrates in-
cluding mud, hard clams prefer a firm bottom. They favor a mixture con-
taining sand or shell which provides points of attachment for juveniles as well
as protection from many predators.
The American oyster in the Chesapeake Bay spawns in the summer when
water temperatures exceed 15 degrees C. Heavy spawning is likely to occur at
22-23 degrees C. Sperm and eggs are released into the water where fertil-
ization occurs, producing free-swimming larvae. The duration of the larval
stage varies with temperature, lasting sometimes as few as 7 to 10 days, but
most often between 2 to 3 weeks before the larvae set and became sessile
organisms. Soft clams and hard clams, like most other bivalve mollusks, spawn
when a critical temperature occurs. In the Chesapeake, soft clams spawn in
the spring when water temperature reaches 10 degrees C and spawning may
be repeated in the fall when water temperature falls to 20 degrees C. Soft clam
eggs develop into planktonic trochophore larvae in about 12 hours. Larvae
remain in the water column for about 6 weeks during the fall. The faster
spring rate of larval development is caused by temperatures at the warmer end
of the soft clam's spawning temperature range. Setting of soft clams, there-
fore, may occur twice in the same year. Frequently, however, heavy predation
on the spring set by blue crabs and bottom-feeding fish results in unsuccess-
ful recruitment. Hard clams spawn at temperatures of 22-24 degrees C. Normal
egg development occurs between 20-35 ppt salinity. At salinities below 17.5
ppt, larvae fail to metamorphose and growth of juveniles ceases. Optimal
temperatures for larval growth range between 18 and 30 degrees C. Growth
ceases at oxygen concentrations below 2.4 mg/1.
TROPHIC IMPORTANCE
The American oyster is an epibenthic suspension feeder, ingesting a
variety of algae, bacteria, and small detrital particles, most within a range of
3-35 um. Capture efficiency decreases rapidly at particle sizes < 3 um. Particles
filtered but not ingested by the oyster are eliminated as pseudofeces. Fecal and
-44-
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pseudofecal material is important in sediment production and deposition,
providing sites for remineralizing bacterial action, and as a food source for
deposit feeders. The hard shell provides a substrate for numerous epifaunal
organisms such as barnacles and mussels. These characteristics make the
oyster an important member of the benthic community throughout the Chesa-
peake Bay. Oysters, especially in the juvenile stages, are subject to heavy
parasitism and predation by many organisms include protozoans, crabs, snails,
and flatworms.
Both soft and hard clams are also important benthic species in the Bay.
Both species are infaunal suspension feeders, ingesting small detrital particles
and phytoplankton, as well as bacteria and microzooplankton in the case of
My a spp. Adult soft clams burrow deeply, feeding through a long extensible
siphon. Juveniles, burrowing less deeply, often fall prey to finfish, blue crabs
and waterfowl. Commercial harvesting of adults reduces adult populations and
exposes juveniles to predation before they can burrow back into the sediment.
Hard clams favor shallow burrows and are also preyed upon by fish, crabs, and
waterfowl, particularly during the juvenile stage. Also of commercial impor-
tance, the hard clam populations in the Bay suffer from irregular recruitment
and are strictly limited to higher salinity regions.
OTHER SENSITIVITIES
Oysters are sensitive to both turbidity and sedimentation. Excessive sedi-
ment deposition smothers adults and prevents setting of spat. The observation
that the upstream limit of producing oyster bars has shifted downstream
several miles in historic times is evidence of the impact of sedimentation.
Adult feeding rates are depressed at suspended solids concentrations above 24
mg/1 and feeding ceases at concentrations above approximately 50 mg/1. Soft
clams are vulnerable to sediment disturbances since they are slow re-
burrowers. As such, they are impacted by harvesting practices, waves,
currents and bioturbation. Regrowth of SAV would benefit these bivalves by
reducing the amount of sediment resuspension and the resulting turbidity.
Areas of good circulation produce better setting and survival of young
oysters. Most oysters in the Chesapeake are found in areas less than 10 m deep
in which circulation patterns promote adequate levels of dissolved oxygen.
Soft clams are also impacted by anoxia which restricts their distribution to
shallow waters less than 10 m in depth.
Oyster diseases, notably Haplosporidium nelsoni ("MSX") and Perkinsus
marinus ("dermo"), have caused significant mortality in the lower Bay. The
organisms causing these diseases require the higher salinities of the lower
Bay to proliferate. The devastating oyster diseases, MSX and dermo, may not be
restricted by salinity. Infection rate may be related to the oyster's cellular
responses to salinity. In the Choptank River, at salinities < 13 ppt, MSX has
been observed.
Temperatures of 32.5 degrees C or greater are lethal to adult soft clam
limiting intertidal distribution in the species' southern range. For oysters,
soft clams, and hard clams, it is generally agreed that food availability is
another significant factor dictating their survival. Foods of critical sizes are
needed for the different life stages; with the cell sizes generally ranging from
3-35 um.
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TARGET SPECIES: Blue crab (Callinectes sapidus)
Critical life stage: larval, pre-molt, post-molt
Critical life period: June - October
BACKGROUND
Blue crabs are found from the mouth of the Bay to tidal fresh areas.
There are distinct differences in the ranges of males and females. During the
summer months, males are found from freshwater to the polyhaline zone,
although they occur in the greatest numbers in salinities of 3-15 ppt. Maxi-
mum numbers of females occur down Bay at salinities of 10 ppt to ocean
salinities. When air temperatures drop below 10 degrees C, adult crabs leave
shallow, inshore waters and seek deeper areas where they bury themselves
and remain in a state of torpor throughout the winter.
SPAWNING AND DISTRIBUTION
After mating, females migrate south in the Bay toward higher salinity
waters. The timing of egg hatching is seasonally dependent. If mating occurs
during spring, then the first egg mass, or sponge, may hatch in July. Most
females mate during the late summer season in July, August, or September,
with hatching delayed until the following summer. A female may also produce
two or more egg sponges later in the summer. Blue crab spawning appears to
be concentrated at the mouth of Chesapeake Bay in the channel region
between Cape Henry and Cape Charles where salinities are near oceanic.
McConaugha et al. (1981) examined seasonal, horizontal and vertical dis-
tribution of blue crab larvae in the mouth of the Chesapeake Bay and
nearshore waters. Only early stage zoeae (mainly Stages I-III) and megalopae
were found in the Bay mouth, while all zoeal stages and megalopae were
present in abundance offshore. They concluded that larval development
occurs in the rich coastal waters and recruitment back to the estuary occurs in
the post-larval or juvenile stages.
Juvenile crab migration up the Chesapeake Bay and its tributaries begins
in August. Male and female juvenile crabs apparently have different migra-
tory patterns. Miller et al. (1975) reported that juvenile crabs, predominantly
males, move into the Chesapeake and Delaware Canal area in late spring. This
distribution of sexes is quite unlike the sex distribution of juvenile crabs in
the lower Bay, around Tangier Sound, suggesting there is a separation of the
sexes at an early stage which is probably due to differences in migratory
behavior.
GROWTH
Blue crab growth is regulated by water temperature. Growth occurs from
late April to mid-October when temperatures are above 15 degrees C (Van
Engel et al., 1973). They grow by shedding their hard shells (molting).
Molting is a major physiological event of crustacean life history. Blue crabs
molt frequently during the early juvenile stages (7-10 days). The periodicity
decreases with age and size. The premolt and postmolt phases are periods of
high metabolic activity; therefore, the animal may be more susceptible to
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environmental stress during these periods. The crabs reach adult size (130 mm
or larger) while on the "nursery grounds," brackish water habitats in the
tributaries and mainstem of the Bay.
TROPHIC IMPORTANCE
Blue crabs are generally considered omnivorous. The zoeae and mega-
lopae prey primarily upon zooplankton. The megalopae will also feed upon
pieces of fish or shellfish and aquatic plants (Van Engel, 1958). Juvenile and
adult blue crabs are also omnivorous, feeding on benthic macroinvertebrates,
small fish, aquatic vegetation and associated fauna, and dead organisms
(Lippson et al., 1979). The blue crab is known to prey on young quahogs and
seed oysters under experimental conditions. It will also prey on oyster spat,
newly set oysters and clams, or young oysters if other food is unavailable (Van
Engel, 1958; Shea et al., 1980). It follows that the blue crab may be a major
factor in the control of benthic populations (Shea et al., 1980).
TARGET SPECIES: Canvasback (Aythya valisineria)
Critical Life Stage: nestling
Critical Life Period: March - June
BACKGROUND
The canvasback is a diving duck, often descending several meters in
search of food. It breeds on the North American prairies and migrates only
when water becomes too cold in its summer range. Chesapeake Bay popu-
lations have been reduced from a peak of almost 400,000 canvasbacks, to aver-
ages of 250,000 in the 1950s and generally less than 70,000 in the 1980s. Before
hunting reforms in 1918, canvasbacks, an international delicacy, were
slaughtered in the thousands by market hunters.
Canvasbacks have adapted with success from their earlier dependence on
and preference for wild celery and other submerged aquatic vegetation. These
ducks now depend on Rangia and Macoma clams, snails, insects, worms and
small crustaceans as a substantial portion of their diet. This dietary change
has made them less desirable table fare, but canvasbacks are still much prized
by hunters.
TARGET SPECIES: Redhead duck (Aythya americana)
Critical Life Stage: nestling
Critical Life Period: March - June
BACKGROUND
The redhead's principal breeding grounds are the North American
prairies, where habitats have been reduced. Most redheads migrate to the Gulf
of Mexico coast, but in the 1950s as many as 118,800 were estimated in the
Chesapeake Bay during January 1956. The 1980s populations have averaged
about 3,500. This duck's exceptionally large salt glands enable it to spend much
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of its wintering time in waters at or near ocean salinity. Entire winters may
be spent on the water.
The food of the redhead consists largely of vegetation, more so than other
diving ducks. Sago pondweed, wild celery, widgeongrass and other submerged
aquatic plants are the favored items. A small percentage of insects, mollusks,
other invertebrates, and small fish are also eaten.
TARGET SPECIES: Black Duck (Anas rubripes)
Critical Life Stage: nestling
Critical Life Period: April - July
BACKGROUND
The Chesapeake Bay's population of black ducks has dwindled in recent
years, from an estimated 200,000 overwintering in 1955 to less than 50,000 in
the mid-1980s. For this reason, more severe hunting restrictions have been
placed upon the species.
Black ducks pair in the autumn. Typically in April, the female lays from
7 to 12 eggs in simple, hollowed-out, pine needle-lined nests. In the Chesa-
peake Bay area, isolated islands and marshes are the favored breeding places.
Though wary of people and other intruders such as predators, which include
raccoons, crows and gulls, almost half the nests are usually destroyed. A
second clutch of eggs is then usually laid.
Black ducks feed on animal foods more than most other dabblers. Favored
items are snails, mussels, clams, small crustaceans and immature insects.
Pondweeds (Potamogeton spp.), widgeongrass, eelgrass, smooth cordgrass, wild
rice and bulrushes are plant food items which, along with corn, are consumed
when available.
TARGET SPECIES: Wood duck (Aix sponsa)
Critical Life Stage: nestling
Critical Life Period: April - July
BACKGROUND
Wood ducks are at the northern edge of their wintering range in the
Chesapeake area, but can breed successfully, given proper habitat. Breeding
habitat should include 10 acres of isolated wetlands with at least 50 percent
cover, while wintering habitats may be less dependent on size given the
adults' greater sociability and mobility. Typical habitat consists of secluded
freshwater swamps and marshes providing plenty of downed or overhanging
trees, shrubs, and flooded woody vegetation. Areas inhabited by beaver often
provide good wood duck habitat. Cavity nesting sites are important for wood
ducks, in order to provide safety from predators such as raccoons.
Adults are largely herbivorous, typically feeding on nuts and fruits from
woody plants, aquatic plants and seeds. Their diet does include some insects
-56-
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and aquatic invertebrates. During the egg laying period, adult wood duck hens
have high protein and calcium requirements, satisfied mainly through an
invertebrate diet. Young ducklings up to 6 weeks of age also ingest a high
percentage of invertebrates, chiefly insects.
TARGET SPECIES: Great blue heron (Ardea herodeas)
Critical Life Stage: nestling
Critical Life Period: May-July
BACKGROUND
Habitat for the great blue heron includes wooded areas suitable for colo-
nial nesting and wetlands within a specified distance (e.g. 1 kilometer) of a
heronry where foraging can occur. The heronry area itself can be an acre or
two in size, but is preferably isolated. Most great blue heron colonies in the
Bay area are located in riparian swamps with trees tall enough for nest place-
ment at 5 to 15 m above ground. Other wading bird species may coexist in a
great blue heronry. Four eggs are typically laid by the adult female, with an
incubation period of four weeks.
Great blue herons feed alone or occasionally in flocks. Feeding usually
occurs during the day, but occasionally takes place at night. Both still-hunt-
ing and stalking techniques are used to hunt for fish which is their main
prey. Herons also eat frogs, lizards, snakes, small birds, mammals, and insects.
Usually, feeding is limited to clear waters less than 0.5 m in depth, with firm
substrate. Contaminants in the food chain have been documented as a prob-
lem, especially dieldrin and DDE and possibly other organochlorines, which
cause eggshell thinning.
TARGET SPECIES: Great (American) egret (Casmerodius albus)
Critical Life Stage: nestling
Critical Life Period: June - August
BACKGROUND
Habitat needs of the great heron are similar to those of the great blue
heron; a heronry area preferably isolated, with good roosting trees and a
foraging area close by. Fresh, brackish and salt water marshes are all used for
foraging.
Three or four eggs, incubating in about 24 days, are typically produced.
The large nests can be from 6 to greater than 15 meters high, located in large
trees near the water. Crows and vultures may prey on the eggs when left
unattended. The young of the year sometimes wander northward before
migrating southward for the winter.
The food of the great egret consists of small fish from the shallow waters,
as well as frogs, lizards, small snakes, crustaceans, mollusks and insects. The
depth of water in which foraging takes place is usually less than 25 cm.
-60-
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itical Life period:
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TARGET SPECIES: Little blue heron (Florida cae.ru.lea)
Critical Life Stage: nestling
Critical Life Period: June - August
BACKGROUND
The little blue heron breeds in the Chesapeake Bay area, but winters to
the south. This heron's habitat includes fresh and salt water marshes where it
seeks to avoid human activity. The heronry is typically situated in dense
vegetation on or near a secluded small water body, often far inland from the
larger marsh.
Food for little blue herons consists of minnows, crustaceans, insects such
as grasshoppers, small frogs, lizards and worms. The little blue heron is an
active feeder. Organochlorine residues have probably found their way into
tissues and eggshells, but resulting physiological problems have not been
noted.
TARGET SPECIES: Green heron (Butorides striatus)
Critical Life Stage: nestling
Critical Life Period: June - August
BACKGROUND
The green heron breeds in the Chesapeake Bay area and winters further
to the south. Habitat for the green heron consists of either fresh or saltwater
marsh. This heron appears to be more tolerant of human activity than some
other heron species. The green heron nests singly or in small colonies, unlike
the large heronries of other species. Their nests are not necessarily located
near the water. Four to five eggs are usually laid, with incubation taking 17
days.
Food of the green heron includes minnows, tadpoles, water insects and
their larvae, and crustaceans. They occasionally feed in the uplands where
prey includes worms, insects such as crickets and grasshoppers, snakes and
small mammals.
TARGET SPECIES: Snowy egret (Egretta thula)
Critical Life Stage: nestling
Critical Life Period: June - August
BACKGROUND
The snowy egret breeds in the Chesapeake Bay area and winters to the
south. Both fresh and saltwater marshes are typical habitats for the snowy
egret. Large rookeries, preferably in isolated sections of a marsh, are favored.
Nests usually range in height from 3 to 6 meters in small trees.
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The snowy egret usually produces 4-5 eggs which incubate in about 18
days. Both parents share in nesting chores. Food consists of small fish,
insects, crayfish, small snakes, frogs and lizards.
TARGET SPECIES: Bald eagle (Haleaeetus leucocephalus)
Critical Life Stage: nestling
Critical Life Period: late-January to mid-June
BACKGROUND
The southern bald eagle is still endangered but has been making a come-
back in the Chesapeake Bay area — it was estimated that 136 pairs occupied
nests in 1986. The bald eagle breeds in the Bay area and a select number
migrate south in autumn. Others remain in congregations in areas such as
Caledon State Park, VA, on the Potomac River.
Habitat for the bald eagle is typically close to the water, where tall trees
provide good perching places for the bird to observe prey. The bald eagle
avoids human activities and it will usually not vigorously defend a nest.
Two to three eggs are produced, laid in a large nest up to 7 feet high by 7
feet across. The nest may be 60 feet or more above ground placed in large
trees. About 35 days are required for incubation of eggs.
Food of bald eagles consists primarily of fish, which is often found dead
by the birds. Other dead animals may also be taken. The bald eagle will also
take other prey alive such as ducks, and small to medium mammals. The prob-
lem of organochlorine pesticide residues which caused eggshells to thin and
hatch success to be reduced has been minimized.
TARGET SPECIES: Osprey (Pandion halaetus)
Critical Life Stage: nestling
Critical Life Period: April to mid-July
BACKGROUND
The Chesapeake Bay region supports over 1,500 nesting pairs of ospreys.
Ospreys always live near the water, roosting in large trees and building large,
bulky, stick-nests in trees or on poles or platforms. The osprey can learn to
tolerate human disturbance near its nest. After the breeding and rearing
season is complete, the birds migrate to tropical wintering grounds.
Ospreys feed almost exclusively on live fish taken from near-surface
waters. Nearly every common Chesapeake Bay species of fish has been
recorded in the osprey's diet. Situated at the top of the food chain, ospreys
experienced trouble with accumulated organochlorine pesticide residues of
DDT and dieldrin some years ago. The problems of thinned eggshells and poor
hatch rates experienced at the time, have apparently been rectified, and the
birds are doing well in the Bay.
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LITERATURE CITED
Abraham, B.J. and P.L. Dillon. 1986. Species profiles: life histories and environmental
requirements of coastal fishes and invertebrates. (Mid Atlantic)--Soft shell clam.
U.S. Fish and Wildl. Serv. FWS/OBS-82/11.68. U.S. Army Corps of Engineers, TR EL-
82-4. 18 pp.
Barnes, R.D. 1974. Invertebrate Zoology. 3rd ed. W.B. Saunders Company. 841 pp.
Bason, W.H. 1971. Ecology and early life history of striped bass, Morone saxatilis, in
the Delaware Estuary. Ichthyol. Assoc. Bull. 4, Ichthyological Associates, Box 35,
R.D. 2, Middletown, DE, 19709, 112 pp.
Beauchamp, R.G. (ed). 1974. Marine Environment Planning Guide for the Hampton
Roads/Norfolk Naval Operating Area. Naval Oceanographic Office. Spec. Pub. No.
250. Naval Ocn. Off. Washington, D.C. 262 pp.
Berggren, T.J. and J.T. Lieberman. 1978. Relative contribution of Hudson, Chesapeake
and Roanoke striped bass, Morone saxatilis, stocks to the Atlantic coast fishery.
Fish. Bull. U.S. 76:335-345.
Bigelow, H.B. and W.C. Schroeder. 1953. Striped Bass. In: Fishes of the Gulf of Maine.
U.S. Fish and Wildl. Serv. Fish Bull. 74(53):389-405.
Bogdanov, A.S., S.I. Doroshev and A.F. Karpevich. 1967. Experimental transportation
of Salmo gairdneri and Roccus saxatilis from the USA for acclimatization in bodies
of water in the USSR. Voprosy Ikhtiologii, 7, No. 1.
Boreman, J. 1981. American shad stocks along the Atlantic coast. National Marine
Fisheries Center, Northeast Fisheries Center Lab. Ref. Doc. No. 81-40.
Boynton, W.R., E.M. Setzler, K.V. Wood, H.H. Zion, and M. Homer. 1977. Final report on
Potomac River fisheries study. Ichthyoplankton and juvenile investigations.
Univ. Maryland Center for Environmental and Estuarine Studies, Chesapeake
Biological Laboratory, Solomons, MD, 20688. 328 pp. UMCEES Ref. No. 77-196 CBL.
Bradford, A.D., J.G. Miller, and K. Buss. 1966. Bioassays on eggs and larval stages of
American shad, Alosa sapidissima.. Pages 52-60 in F.T. Carlson, 1968, Suitability of
Interior Maryland Board of Natural Resources, New York Conservation
Department, and Pennsylvania Fish Commission, Washington, D.C. 60 pp.
Brousseau, D.J. 1978. Spawning cycle, fecundity and recruitment in a population of
soft-shell clams, Mya arenaria, from Cape Ann, Massachusetts. Fish. Bull.
76(1):155-166.
Boynton, W.R., T.T. Polgar, and H.H. Zion. 1981. Importance of juvenile striped bass
food habits in the Potomac estuary. Trans. Am. Fish. Soc. 110:56-63.
-72-
-------
Butler, P.A. 1963. Commercial fisheries investigations, pesticide-wildlife studies:
A review of Fish and Wildlife Service investigations during 1961-1962. U.S. Dept.
Inter. Fish Wildl. Circ. 167:11-25.
Butler, P.A. 1964. Pesticide-wildlife studies, 1963. A review of fish and wildlife service
investigations during the calendar year. U.S. Dept. Inter. Fish Wildl. Circ. 199.
Butler, P.A. 1965. Commercial fisheries investigations. Effects of pesticides on fish
and wildlife, 1964. Research findings Fish Wildl. Serv. U.S. Inter. Fish Wildl. Circ.
Butler, P.A. 1966. The problem of pesticides in estuaries. Trans. Am. Fish. Soc. Spec.
Publ. 3:110-115.
Butler, P.A., A.J. Wilson and AJ. Rick. 1960. Effect of pesticides on oysters.
Proc. Nat. Shellfish Assn. 51:23-32.
Calabrese, A., R.S. Collier, D.A. Nelson and J.R. Maclnnes. 1973. The toxicity of heavy
metals to embryos of the American oyster Crassostrea virginica.. Mar. Biol. 18:162-
166.
Calabrese, A. and D.A. Nelson. 1974. Inhibition of embryonic development of the hard
clam, Mercenaria mercenaria by heavy metals. Bull. Environ. Contam. Toxicol.
11:92-97.
Calabrese, A., J.R. Maclnnes, D.A. Nelson and I.E. Miller. 1977. Survival and growth of
bivalve larvae under heavy metal stress. Mar. Biol. 41:179-184.
Cardwell, R.D., D.G. Foremen, T.R. Payne, and DJ. Wilbur. 1976. Acute toxicity of
selected toxicants to six species of fish. Ecol. Res. Ser. EPA 600/3-76-008. Environ.
Res. Lab., U.S. Environ. Prot. Agency, Duluth, MN. 117 p.
Carlson, F.T. 1968. Suitability of the Susquehanna River for the restoration of shad.
U.S. Department of the Interior, Maryland Board of Natural Resources, New York
Conservation Department, and Pennsylvania Fish Commission, Washington, D.C. 60
pp.
Carriker, M.R. 1961. Interrelation of functional morphology, behaviour, and
autecology in early stages of the bivalve Mercenaria mercenaria. J. Elisha
Mitchell Sci. Soc. 77:168-241.
Chittenden, M.E., Jr. 1969. Life history and ecology of the American shad, Alosa
sapidissima, in the Delaware River. Ph.D. Thesis. Rutgers University, New
Brunswick, N.J. 458 pp.
Chittenden, M.E., Jr. and J.R. Westman. 1967. Highlights of the American shad on the
Delaware River. Dept. Environ. Sci. Rutgers Univ. 9 pp.
Colton, J.B., Jr., W.G. Smith, A.W. Kendall, Jr., P.L. Berrien, and M.P. Fahay. 1979.
Principal spawning areas and times of marine fishes, Cape Sable to Cape Hatteras.
U.S. Natl. Mar. Fish. Serv. Fish. Bull. 76:911-914.
Cooper, J.C. and T.T. Polgar. 1981. Recognition of year-class dominance in striped bass
management. Trans. Am. Fish. Soc. 110:180-187.
-73-
-------
Connery, J. (ed.) 1987. " Report of the Workshop on Habitat Requirements for
Chesapeake Bay Living Resources". Eastern Research Group, Arlington, MA.
Costlow, J.D., Jr. 1967. The effect of salinity and temperature on survival and
metamorphosis of megalops of the blue crab Callinectes sapidus Helgolander wiss.
Meeresunters. 15:84-97.
Costlow, J.D., Jr., and C.G. Bookhout. 1959. The larval development of Callinectes
sapidus Rathbun reared in the laboratory. Biol. Bull. 116:373-396.
Crisp, D.J. 1967. Chemical factors inducing settlement in Crassostrea virginica
(Gmelin). J. Anim. Ecol. 36:329-335.
Daniel, D.A. 1967. A laboratory study to define the relationahip between survival of
young striped bass (Morone saxatilis) and their food supply. State of California,
The Resources Agency. Dept. Fish and Game, Anadromous Fisheries Branch,
Administrative Rept. No. 76-1. 13 pp.
Davis, J. 1973. Spawning sites and nurseries of fishes of the genus Alosa in Virginia.
Pages 140-141 in A.L. Pacheco, ed. Proceedings of a workshop on egg, larval and
juvenile stages of fish in Atlantic Coast estuaries. Tech. Publ. No. 1, Natl. Mar. Fish.
Serv. Mid. Atlantic Coast. Fish. Cent., Highlands, N.J. 338 pp.
Davies, W.P. 1970. The effect of temperature, pH and total dissolved solids on the
survival of immature striped bass, Morone saxatilis (Walbaum). PhD. Thesis, North
Carolina State Univ., Raleigh, 100 pp.
Davis, H.C. 1953. On food and feeding of larvae of the American oyster
C. virginica. Biol. Bull. 104:334-350.
Davis, H.C. and A. Calabrese. 1964. Combined effects of temperature and salinity on
development of eggs and growth of larvae of M. mercenaria and C. virginica. Fish.
Bull. 63:643-655.
Doroshev, S.I. 1970. Biological features of the eggs, larvae, and young of the striped
bass, Roccus saxatilis (Walbaum), in connection with the problem of its
acclimatization in the U.S.S.R. J. Ichthyol. 10:235-248.
Dovel, W. L. 1971. Fish eggs and larvae of the upper Chesapeake Bay. Natural
Resources Inst., U. of Md., Spec. Rept. (4): iii+ 71.
Durbin, A.G. 1976. The Role of Fish Migration in Two Coastal Ecosystems: The Atlantic
menhaden (Brevoortia tyrannus) in Narragansett Bay, and the alewife (Alosa
pseudoharengus) in Rhode Island ponds. PhD. Disseration. Univ. of Rhode Island,
Kingston, RI.
Fay, C.W., RJ. Neves, and G.B. Pardue. 1983. Species profiles: life histories and
environmental requirements of coastal fishes and invertebrates (mid-Atlantic) -
Striped bass. U.S. Fish and Wildlife Service. WWS/OBS-82/11.8.
Galtsoff, P.S. 1964. The American oyster, Crassostrea virginica Gmelin. Fish. Bull.
64:1-480.
-74-
-------
Grant, G.C. and J. E. Olney. 1982. Assessment of larval striped bass, Moronejaxatilis
(Walbaum), stocks in Maryland and Virginia waters. Part II. Assessment of
Spawning Activitity in Major Virginia Rivers. Final report, Segment 2, to the Nat'l
Marine Fisheries Service, Gloucester, MA, (Grant No. NA81FAD - VA3B), 42 pp.
Hardy, J. D, Jr. 1978. Development of fishes of the mid-Atlantic Bight: an atlas of the
egg, larval and juvenile stages, Volume III. U.S. Fish. Wildl. Serv., Biol. Serv. Prog.
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Finfish:
References for Key Species of Finfish cited in the Habitat
Requirements Matrices
Habitat Suitability Index Models:
(1) Striped bass FWS/OBS 82/11.8. 1983. 27 pp.
FWS/OBS 82/10.1 1982 23 pp.
FAO Synopsis No. 121- 1980 6/ pp.
(2) Blueback herring FWS/OBS 82/11.9 1983 20pp.
Alewife FWS/OBS 82/10.58 1983 17 pp.
(3) American shad Biological Report 82(10.88) 1985 27 pp.
Hickory shad Biological Report 82(11.45) 1986 15 pp.
Biological Report 82(11.37) 1985 15 pp.
(4) Yellow perch FWS/OBS 82/10.55 1983 32 pp
(5) White perch FWS/OBS 82/.11.7 1983 10pp.
(6) Menhaden FWS/OBS 82/11.11 1983. 15 pp.
(7) Spot FWS/OBS 82/10.20 1982 10 pp.
* All the above publications are from the U.S. Fish and Wildlife Service.
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These sources supplied most of the life history information quoted; additional
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cited above.
Shellfish:
References for Key Species of Shellfish Cited in the Habitat
Requirements Matrices
Habitat Suitability Index Models:
American oyster Biological Report 82(11.65) FWS 1986. 17 pp.
Blue crab FWS/OBS - 82/11.19 1984 13 pp.
Soft shell clam Biological Report 82(11.68) FWS 1986 15 pp.
Hard shell clam Kaumeyer and Setzler-Hamilton. 1982.
* These sources supplied most of the life history information quoted;
additional information on food, contaminants, etc. was taken from
the more general sources cited above.
Waterfowl:
References for Key Species of Birds Cited in the Habitat Requirements Matrices
Habitat Suitability Index Models:
(1) Wood Duck FWS/OBS 82/10.43. 1983. 27 pp.
(2) Redhead (wintering) FWS/OBS 82/10.53. 1983. 14 pp.
(3) American black duck (wintering) FWS/OBS 82/10.68 1984. 16 pp.
* All the above publications are from the U.S. Fish and Wildlife Service,
U.S. Dept. of Interior, Wash. DC 20240.
Bent, A.C. 1962. Life Histories of North American Wildfowl, Part. 1 Dover Publications,
Inc., New York, NY. 239 pp.
Johngard, P.A. 1975. Waterfowl of North America. Indiana University Press. 575 pp.
-84-
-------
These sources supplied most of the life history information quoted; additional
information on food, contaminants, etc. was taken from the more general sources
cited below.
Wading Birds:
Habitat Suitability Index Models:
(1) Great blue heron FWS/OBS 82(10.99). 1985. 23 pp.
(2) Great egret FWS/OBS 827(10.78). 1984. 23 pp.
The above publications are from the USFWS, U.S. Dept of Interior, Wash. DC 20240.
Bent, A.C. 1963. Life Histories of North American Marsh Birds. Dover Publications,
Inc., New York, NY. 385 pp.
Erwin, R.M. 1979. Coastal Waterbird Colonies: Cape Elizabeth, Maine to Virginia.
FWS/OBS-79/10. 212 pp.
See also general references below.
Raptors:
Bent, A.C. 1961. Life Histories of North American Birds of Prey. Part 1. Dover
Publications, Inc., New York, NY. 398 pp.
Bird, D.M., N.R. Seymour and J.M. Gerrard. 1983. Biology and Management of Bald
Eagles and Ospreys. MacDonald Raptor Research Center of McGill University -
Proceedings of 1st International Symposium, Montreal, Canada, October 1981.
325 pp.
U.S. Fish and Wildlife Service. 1982. The Chesapeake Bay Region Eagle Recovery Plan.
Region 5, USFWS. 81 pp.
General:
Fish and Wildlife Service. 1951. Food of Game Ducks in the United States and Canada.
Research Report 30. U.S. Dept. of Interior. 308 pp.
Martin, A.C., H.S. Zim and A.L. Nelson. 1961. American Wildlife and Plants - A Guide to
Wildlife Food Habits. Dover Publications, Inc., New York, NY. 500 pp.
Collins, H.H., Jr., Ed. 1981. Complete Field Guide to North American Wildlife. Eastern
Edition. Harper and Row, Publishers, New York. 714 pp.
Wernert, S.J., Ed. 1982. North American Wildlife. Readers Digest Association, Inc.,
Pleasantsville, NY. 539 pp.
Stevenson, J.C. and N. Confer. 1978. Summary of Available Information on Chesapeake
Bay Submerged Vegetation. FWS/OBS 78/66. U.S. Fish and Wildlife Service, co-
sponsored by Maryland Dept. of Natural Resources and U.S. Environmental
Protection Agency. 335 pp.
-85-
-------
Contaminant Sources:
U.S. EPA. 1982. Chesapeake Bay Program Technical Studies: A Synthesis. Part IV
SAV. pp. 379-634.
Brown, A.W.A. 1978. Ecology of Pesticides. John Wiley & Sons, Inc., New York.
525 pp.
Ohlendorf, H.M., E.E. Klaas and T.E. Kaiser. 1979. Environmental Pollutants and
Eggshell Thickness: Anhingas and Wading Birds in the Eastern U.S. Special
Scientific Report - Wildlife #216. USFWS, U.S. Dept. Of Interior. 94 pp.
* Provided by U.S. Fish and Wildlife Service. (1987)
-86-
-------
APPENDIX A:
TOXICITY OF SUBSTANCES TO STRIPED BASS LARVAE AND JUVENILES
Adapted from Westin and Rogers. 1978.
Synopsis of Biological Data on the
Striped Bass, Morone saxatilis
(Walbaum) 1972. University of
Rhode Island, Marine Technical
Report 67, Kingston, RI
-------
-TABLE 1-
TOXICITY OF SUBSTANCES TO STRIPED BASS LARVA
SUBSTANCE
96-HOUR TLm
(95% C.I.)
(mg/1)
AUTHOR
Acriflavine
Aldrin
Ami fur
Butyl ester of 2,4-D
Cadmium
Chloride
Chlorine
Copper
Copper
Copper sulfate
Dieldrin
Diquat
Diuron
Dylox
Ethyl parathion
Formaldehyde
HTH
Iron
Karmex
Malachita green
Methylene blue
Methyl parathion
Potassium dichromate
Potassium permanganate
Roccal
Rotenone
Sulfate
Tad-Tox
Terramycin
Zinc
Zinc
5.0 (NA)
0.01 (NA)
10.0 (NA)
0.15 (NA)
0.001 (NA)
1000 (NA)
0.20 (NA)
0.40-0.07 incipient
0.05 (NA)
0.31 (0.12-3.08)
0.1 (NA)
0.001 (NA)
1.0 (NA)
0.5(NA)
5.0 (NA)
2.0 (NA)
10.0 (NA)
0.5 (NA)
4.0 (NA)
0.5 (NA)
0.05 (NA)
1.0 (NA)
5.0 (NA)
100 (NA)
1.0 (NA)
0.5 (NA)
0.001 (NA)
250 (NA)
5.0 (NA)
50.0 (NA)
0.1 (NA)
1.18 (0.25-2.46)
Hughes (1973)
Hughes (1973)
Hughes (1973)
Hughes (1971)
Hughes (1973)
Hughes (1973)
Morgan & Prince (1977)
Middaugh et al. (1977)
Hughes (1973)
O'Rear (1971)
Hughes (1971)
Hughes (1973)
Hughes (1973)
Hughes (1973)
Hughes (1971)
Hughes (1971)
Hughes (1973)
Hughes (1971)
Hughes (1973)
Hughes (1971)
Hughes (1973)
Hughes (1973)
Hughes (1971)
Hughes (1971)
Hughes (1971)
Hughes (1973)
Hughes (1973)
Hughes (1973)
Hughes (1973)
Hughes (1973)
Hughes (1973)
O'Rear (1971)
a All 4-7 day-old larvae from Moncks Corner, South Carolina, tested at 21
C, except O'Rear (1971) which were tested in 14-19 C range, Morgan &
Prince (1977) not specified, and Middaugh et al. (1977) at 18 C.
b NA = not available (i.e., neither given nor calculatable).
c 48-hour TLm
d 96-hour LCo
e 24-hour TLm
-------
-TABLE 2-
TOXICITY OF SUBSTANCES TO JUVENILE STRIPED BASS
SUBSTANCE
TEST
TEMP C
96-HOUR TLm
(95* C.I.)
(mg/1)
AUTHOR
Abate
Achromycin
Acriflavine
Aldrin
Amifur
Ammonium hydroxide
Aquathol
Bayluscide
Benzene
Butyl ester of
2,4-D
Cadmium
Carbaryl
Casoron
Chlordane
Chloride
Chlorine
Cooling Tover
Slowdown and
Power Plant
Chemical Discharge
Co-Ral
Copper
Copper sulfate
Cutrine
ODD
DDT
Dibrom
Dieldrin
Diquat
Diuron (Karmex)
13 1.0 (NA)
21-22 190 (153.2-235.6)
21 27.5 (NA)
16.0 (14.7-17.4)
13 0.0072 (0.0034-0.0152
21 LCo 0.075 (NA)
20 0.010 (NA)
21 LCo 30.0 (NA)
15 1.9-2.85
23 1.4-2.8
21 610 (634-795)
21 72 hr. 1.05 (0.94-1.18)
17.4 10.9 ul/1 (+0.02)
16 5.8 ul/1
21 3.0 (NA)
20 70.0 (NA)
21 0.002 (NA)
17 1.0 (NA)
21 6,2000 (5,210-7,378)
15 0.0118 (0.0057-0.024)
21 5000 (NA)
18 0.04 incipient
4.5-6.0 >4.0X
18.5-26.0 >4.0X [incipient LC50
w/o CL2, 3.6X
(3.81X -3.4X)]
21 62 (53-73)
21 0.05 (NA)
17 4.3 (NA)
21 0.15 (NA)
21-22 0.6 (0.51-0.83)
21 0.62 (0.54-0.71)
21 0.1 (NA)
17 0.0025 (0.0016-0.004)
17 0.00053 (0.00038-
0.00084)
13 0.5 (0.1-2.4)
14 0.0197 (0.0098-
0.00334)
21 0.25 (NA)
21 10.0 (NA)
21 80 (74-86)
21 6.0 (NA)
Korn & Earnest (1974)
Kelley (1969)
Hughes (1973)
Wellborn (1971)
Korn & Earnest (1974)
Hughes (1973)
Rehwoldt et al. (1977)
Hughes (1973)
Hazel et al. (1971)
ii it n ii
Wellborn (1971)
Wellborn (1971)
Meyerhoff (1975)
Benville and Korn (1977)
Hughes (1971)
Rehwoldt et al. (1977)
Hughes (1973)
Korn & Earnest (1974)
Wellborn (1971)
Korn & Earnest (1974)
Hughes (1973)
Middaugh et al. (1977)
Texas Instruments (1974)
Wellborn (1971)
Hughes (1973)
Rehwoldt et al. (1971)
Hughes (1971)
Kelley (1969)
Wellborn (1969)
Hughes (1973)
Korn & Earnest (1974)
Korn & Earnest (1974)
Korn & Earnest (1974)
Korn & Earnest (1974)
Hughes (1973)
Hughes (1973)
Wellborn (1969)
Hughes (1973)
-------
-TABLE 2 (cont.)-
SUBSTANCE
TEST
TEMP C
96-HOUR TLm
(95% C.I.)
(mg/1)
AUTHOR
Dursban
Dylox
Endosulfan
Endrin
E.P.N.
Ethyl parathion
Fenthion
Formaldehyde
Heptachlor
HTH
Instant Sea
as (Cl)
Iron
Karmex (Diuron)
Lindane
Malachite green
Malathion
Methoxychlor
Methylene blue
Methyl parathion
MS-222
MS-222
with 20 o/oo
Nickel
Oil field brine
(as Cl)
Potassium
dichromate
Potassium
permanganate
13 0.00058 (0.00035-
0.00097)
21 2.0 (NA)
5.2 (4.2-8.0)
16 0.0001 (0.000048-
0.00021)
17 0.000094 (0.000045-
0.00019)
18 0.60 (0.025-0.150)
21 1.0 (NA)
15 0.0178 (0.0048-
0.0657)
13 0.453 (0.216-0.955)
21 15 (NA)
21-22 20 (15.4-26)
21 18 (10-32)
13 0.003 (0.001-0.006)
21 0.25 (NA)
21 LCo 17000 (NA)
21 6.0 (NA)
21 6.0 (NA)
3.1 (2.5-3.9)
21 0.40 (0.35-0.46)
13 0.0073 (0.0045-0.0119)
21 0.2 (NA)
24 hr. 0.30 (0.27-0.33)
21 0.24 (0.20-0.29)
13 0.014 (0.013-0.015)
20 0.039 (NA)
15 0.0033 (0.0021-0.0051)
21 12.0 (NA)
21 4.5 (NA)
13 0.79 (0.17-1.40)
20 14.0 (NA)
21-22 31.5 (25.6-37.5)
22-28 24 hr. 50.0 (NA)
21-22 31.5 (26.6-37.5)
17 6.2 (NA)
21 LCo 16600 (NA)
21 75 (NA)
21 4.0 (NA)
21-22 2.6 (2.17-3.12)
Korn & Earnest (1974)
Hughes (1971)
Wellborn (1969)
Korn & Earnest (1974)
Korn & Earnest (1974)
Korn & Earnest (1974)
Hughes (1971)
Korn & Earnest (1974)
Korn & Earnest (1974)
Hughes (1973)
Kelley (1969)
Wellborn (1969)
Korn & Earnest (1974)
Hughes (1971)
Hughes (1973)
Hughes (1973)
Hughes (1971)
Wellborn (1969)
Wellborn (1971)
Korn & Earnest (1974)
Hughes (1973)
Wellborn (1971)
Wellborn (1971)
Korn & Earnest (1974)
Rehwoldt et al. (1977)
Korn & Earnest (1974)
Hughes (1973)
Hughes (1971)
Korn & Earnest (1974)
Rehwoldt et al. (1977)
Kelley (1969)
Tatum et al. (1965)
Kelley (1969)
Rehwoldt et al. (1971)
Hughes (1968)
Hughes (1971)
Hughes (1971)
Kelley (1969)
-------
-TABLE 2 (cont.)-
SUBSTANCE
Polyotic
PMA
Quinaldine
Quinaldine with
20 o/oo
Reconstituted
sea water
Roccal
Rotenone
Simazine
Sodium nitrilo-
triacetic acid
Sulfate
Syndet Ch
Syndet Ga
Tad-Tox
Terramycin
Toluene
Toxaphene
m-xylene
Zinc
2, 4, 5, T
TEST
TEMP C
21
21
21-22
21-22
22-28
21-22
21-22
21
21
21
20
21
20
21
21
21-22
21
16
17
16
21
17
20
96-HOUR TLm
(95% C.I.)
(mg/1)
2.5 (2.1-2.9)
>1818 (NA)
1.1 (0.84-1.44)
4.5 (3.82-5.45)
24 hr. 22.0 (NA)
5.0 (3.86-6.65)
35 o/oo (NA)
1.5 (NA)
LCo 0.001 (NA)
0.25 (0.17-0.36)
5500 (NA)
3500 (NA)
4.6 (NA)
8.7 (NA)
10.0 (NA)
75.0 (NA)
170 (140.5-205.7)
178 (144-221)
165 (147-185)
7.3 ul/1
0.0044 (0.002-0.009)
9.2 (8.3-10) ul/1
0.1 (NA)
6.7 (NA)
14.6 (NA)
AUTHOR
Wellborn (1969)
Wellborn (1969)
Kelley (1969)
Kelley (1969)
Tatum et al. (1965)
Kelley (1969)
Kelley (1969)
Hughes (1973)
Hughes (1973)
Wellborn (1969)
Eisler et al.
(1972)
Hughes (1973)
Eisler et al.
(1972)
Eisler et al.
(1972)
Hughes (1973)
Hughes (1973)
Kelley (1969)
Wellborn (1969)
Wellborn (1971)
Benville & Korn (1977)
Korn & Earnest (1974)
Benville & Korn (1977)
Hughes (1973)
Rehwoldt et al. (1971)
Rehwoldt et al. (1977)
a Unless specified otherwise
b NA = not available (i.e., neither given nor calculatable)
c Range of 96-hour TLm in freshwater, 33% sea water, and sea water (95%
C.I. given for percent mortality at 0, 40, 60, 80, and 100%).
-------
APPENDIX B:
HABITAT DISTRIBUTION MAPS OP CRITICAL LIFE STAGES OF
THE TARGET CHESAPEAKE BAY LIVING RESOURCE SPECIES
-------
List of Habitat Distribution Haps for the Critical Life Stages of the
Target Chesapeake Bay Living Resource Species
1. 1986 Distribution of Submerged Aquatic Vegetation in Chesapeake Bay
2. Striped Bass (Morone saxatilis): Habitat Distribution of Legislatively
Defined Spawning Reaches and Rivers in Chesapeake Bay
3. Blueback Herring (Alosa aestivalis): Habitat Distribution of Nursery
Areas in Chesapeake Bay
4. Alewife (Alosa pseudoharengus): Habitat Distribution of Nursery Areas
in Chesapeake Bay
5. American Shad (Alosa sapidissima): Habitat Distribution of Nursery
Areas in Chesapeake Bay
6. Hickory Shad (Alosa mediocris): Habitat Distribution of Nursery Areas
in Chesapeake Bay
7. Yellow Perch (Perca flavescens): Habitat Distribution of Spawning
Areas in Chesapeake Bay
8. White Perch (Morone americana): Habitat Distribution of Spawning and
Nursery Areas in Chesapeake Bay
9. Menhaden (Brevoortia tyrannus): Habitat Distribution of Nursery Areas
in Chesapeake Bay
10. Spot (Leiostomus xanthurus): Habitat Distribution of Nursery Areas in
Chesapeake Bay
11. Bay Anchovy (Anchoa mitchelli): Habitat Distribution of Spawning and
Nursery Areas in Chesapeake Bay
12. American Oyster (Crassostrea virginica): Habitat Distribution of Seed
Areas and Suitable Substrate in Chesapeake Bay
13. Softshell Clam (Mya arenaria): Habitat Distribution in Chesapeake Bay
14. Hard Clam (Mercenaria mercenaria): Habitat Distribution in Chesapeake
Bay
15. Blue Crab (Callinectes sapidius): Summer Habitat Distribution of
Females and Spawning Areas in Chesapeake Bay
16. Blue Crab (Callinectes sapidius): Summer Habitat Distribution of Males
in Chesapeake Bay
17. Blue Crab (Callinectes sapidius): Winter Habitat Distribution of
Females in Chesapeake Bay
18. Blue Crab (Callinectes sapidius): Winter Habitat Distribution of Males
in Chesapeake Bay
19. Canvasback (Aythya valisneria): Distribution of Wintering Populations
20. Redhead Duck (Aythya americana): Distribution of Wintering Populations
21. Black Duck (Anas rubripes): Distribution of Wintering Populations
22. Wood Duck (Aix sponsa): Distribution of Wintering Populations
23. Colonial Waterbirds: Habitat Distribution of Nesting Populations in
Chesapeake Bay
24. Osprey (Pandion halaetus) and Bald Eagle (Haliaeetus leucocephalus):
Habitat Distribution of Nesting Populations in Chesapeake Bay
-------
1986 DISTRIBUTION OF SUBMERGED AQUATIC VEGETATION
IN CHESAPEAKE BAY
LEGEND
SAV DISTRIBUTION
SCALE 1:1,500.000
SOURCE: Orth et al.. 1987
FIGURE 1
-------
STRIPED BASS {Morone saxatilis): HABITAT DISTRIBUTION OF
LEGISLATIVELY DEFINED SPAWNING REACHES AND RIVERS
LEGEND
SPAWNING REACHES
SPAWNING RIVERS
SCALE 1:1.500,000
SOURCES: Code of Maryland Regulations 08.02.05.02
Virginia Marine Resources Commission Regulation 450-01-0034
FIGURE 2
-------
BLUEBACK HERRING (Alosa aestivalis): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS:
EGO AND LARVAL STAGES
SCALE 1:1,500.000
SOURCE: Corps of Engineers, 1980
FIGURE 3
-------
ALEWIFE (Alosa pseudoharengus): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS:
EGG AND LARVAL STAGES
SCALE 1:1.500,000
SOURCE: Corps of Engineers, 1980
FIGURE 4
-------
AMERICAN SHAD (Alosa sapidissima): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS:
EGG AND LARVAL STAGES
SCALE 1:1.500,000
SOURCE: Corps of Engineers, 1980
FIGURE 5
-------
HICKORY SHAD (Alosa mediocris): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS:
EGG AND LARVAL STAGES
SCALE 1:1.500,000
SOURCE: Corps of Engineers, 1980
FIGURE 6
-------
YELLOW PERCH (Perca flavescens): HABITAT DISTRIBUTION OF
SPAWNING AREAS IN CHESAPEAKE BAY
LEGEND
SPAWNING AREAS
SCALE 1:1. 500,000
SOURCE: Corps of Engineers, 1980
FIGURE 7
-------
WHITE PERCH (Morone americana): HABITAT DISTRIBUTION OF
SPAWNING AND NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
SPAWNING AREAS
NURSERY AREAS
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 8
-------
MENHADEN (Brevoortia tyrannus): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 9
-------
SPOT (Leiostomus xanthurus): HABITAT DISTRIBUTION OF
NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
NURSERY AREAS
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 10
-------
BAY ANCHOVY (Anchoa mitchilli): HABITAT DISTRIBUTION OF
SPAWNING AND NURSERY AREAS IN CHESAPEAKE BAY
LEGEND
SPAWNING AREAS
NURSERY AREAS
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 11
-------
AMERICAN OYSTER (Crassostrea virginica): HABITAT
DISTRIBUTION OF SEED AREAS AND SUITABLE SUBSTRATE IN
CHESAPEAKE BAY
LEGEND
SEED AREAS
SUITABLE SUBSTRATE
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 12
-------
SOFTSHELL CLAM (Mya arenaria): HABITAT DISTRIBUTION
IN CHESAPEAKE BAY
LEGEND
HIGH DENSITY
LOW DENSITY
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 13
-------
HARD CLAM (Mercenaria mercenaria): HABITAT DISTRIBUTION
IN CHESAPEAKE BAY
LEGEND
HIGH DENSITY
LOW DENSITY
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 14
-------
BLUE CRAB (Callinectes sapidus) : SUMMER HABITAT
DISTRIBUTION OF FEMALES AND SPAWNING AREAS IN CHESAPEAKE
BAY
LEGEND
SPAWNING AREAS
HIGH DENSITY
LOW DENSITY
SCALE 1 1,500.000
SOURCE: Corps of Engineers, 1980
FIGURE 15
-------
BLUE CRAB (Callinectes sapidus) : SUMMER HABITAT
DISTRIBUTION OF MALES IN CHESAPEAKE BAY
LEGEND
HIGH DENSITY
LOW DENSITY
SCALE 1:1,500,000
SOURCE: Corps of Engineers, 1980
FIGURE 16
-------
BLUE CRAB (Callinectes sapidus) : WINTER HABITAT
DISTRIBUTION OF FEMALES IN CHESAPEAKE BAY
LEGEND
HIGH DENSITY
LOW DENSITY
SCALE 1:1.500,000
SOURCE: Corps of Engineers, 1980
FIGURE 17
-------
BLUE CRAB (Callinectes sapidus) : WINTER HABITAT
DISTRIBUTION OF MALES IN CHESAPEAKE BAY
LEGEND
HIGH DENSITY
LOW DENSITY
SCALE 1:1,500.000
SOURCE: Corps of Engineers, 1980
FIGURE 18
-------
CANVASBACK (Aythya valisneria) : DISTRIBUTION OF
WINTERING POPULATIONS IN CHESAPEAKE BAY
LEGEND
WINTERING POPULATIONS
SCALE 1:1,500,000
SOURCE: USFWS unpublished data FIGURE 19
-------
REDHEAD DUCK (Aythya americana) : DISTRIBUTION OF
WINTERING POPULATIONS IN CHESAPEAKE BAY
LEGEND
WINTERING POPULATIONS
SCALE 1:1.500,000
SOURCE: USFWS unpublished data
FIGURE 20
-------
BLACK DUCK (Anas rubripes) : DISTRIBUTION OF
WINTERING POPULATIONS IN CHESAPEAKE BAY
LEGEND
WINTERING POPULATIONS
SCALE 1:1.500.000
SOURCE: USFWS unpublished data
FIGURE 21
-------
WOOD DUCK (Aix sponsa) : DISTRIBUTION OF
WINTERING POPULATIONS IN CHESAPEAKE BAY
LEGEND
WINTERING POPULATIONS
SCALE 1:1,500,000
SOURCE: USFWS unpublished data
FIGURE 22
-------
COLONIAL WATERBIRDS: HABITAT DISTRIBUTION OF NESTING
POPULATIONS IN CHESAPEAKE BAY
LEGEND
NESTING POPULATIONS
SCALE 1:1. 500,000
w
NOTE: Colonial waterbirds include: Great blue heron (Ardea herodias);
Little blue heron (Florida caeru/ea); Green-backed heron (Butorides striatus);
Snowy egret (Egretta thula); American or great egret (Casmerodius albus)
Scattered nests may occur in many other wooded, secluded areas of Bay tributaries.
SOURCE: USFWS unpublished data FIGURE 23
-------
OSPREY (Pandion haliaetus) AND BALD EAGLE (Haliaeetus
leucocephalus): HABITAT DISTRIBUTION OF NESTING POPULATIONS
IN CHESAPEAKE BAY
LEGEND
1"^| NESTING POPULATIONS
SCALE 1:1,500,000
NOTE: Bald eagle nests, roosts and feeding areas are generally found within one mile of
the riverine and estuarine shoreline in the Bay system. Occasionally, lakes and
reservoirs are used. Some bald eagles remain in the Bay area year round.
SOURCE: USFWS unpublished data
FIGURE 24
-------
APPENDIX C:
REPORT ON THE WORKSHOP ON HABITAT REQUIREMENTS FOR
CHESAPEAKE BAT LIVING RESOURCES
-------
United States Environmental Protection Agency
CBP/TRS 8/87
July 1987
Report of the Workshop on
Habitat Requirements for
Chesapeake Bay Living Resources
m ^^»-
Chesapeake
Bay
Program
-------
REPORT OF THE WORKSHOP ON
HABITAT REQUIREMENTS FOR
CHESAPEAKE BAY LIVING RESOURCES
Annapolis, Maryland
February 24, 1987
Prepared by:
Jan Connery
Eastern Research Group, Inc,
6 Whittemore Street
Arlington, MA 02174
Submitted to:
Chesapeake Bay Program's
Living Resources Task Force
FINAL REPORT
May 29, 1987
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TABLE OF CONTENTS
1.
2.
3.
4.
5.
PREFACE
BENTHOS PLANNING SESSION
PLANKTON PLANNING. SESSION
2.1
2.2
2.3
Introduction
Hypotheses
Conclusion
SUBMERGED AQUATIC VEGETATION PLANNING SESSION
3.1
3.2
Introduction
3.1.1 High Salinity
3.1.2 Mesohaline
3.1.3 Freshwater
General Comments/Recommendations
SHELLFISH PLANNING SESSION AND TECHNICAL WORK GROUP
4.1
4.2
4.3
4.4
General Approach and Recommendations
Geographic Distribution
Critical Life Stage and Period
Habitat Requirements
FINFISH PLANNING SESSION AND TECHNICAL WORK GROUP
5.1
5.2
5.3
Species List
General Changes and Recommendations
Matrices
5.3.1 Striped Bass
5.3.2 Alewife/Blueback Herring
5.3.3 Atlantic Menhaden and Spot
5.3.4 Other Species
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4
4
5
6
8
8
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10
12
12
13
13
14
20
20
22
28
28
34
37
40
5.4 Conclusions 40
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TABLE OF CONTENTS (CONT.)
Page
6. WATERFOWL/BIRDS PLANNING SESSION AND TECHNICAL 41
WORK GROUP
6.1 Approach 41
6.2 General Changes and Recommendations 41
6.3 Waterfowl 42
6.3.1 Canvasback 43
6.3.2 Redhead 47
6.3.3 Black Duck 48
6.3.4 Wood Duck 49
6.4 Wading Birds 51
6.4.1 Great Blue Heron 51
6.4.2 Little Blue Heron and 53
Green-Backed Heron
6.5 Raptors 53
6.5.1 Osprey 53
6.5.2 Bald Eagle 54
7. BLUE CRAB TECHNICAL WORK GROUP 56
7.1 Introduction 56
7.2 Critical Life Stage and Period 56
7.3 Background 56
7.4 Matrix 57
7.5 Geographic Distribution 60
8. REFERENCES 61
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APPENDIX A
APPENDIX B
APPENDIX C
APPENDIX D
APPENDIX E
APPENDIX F
TABLE OF CONTENTS (CONT.)
WORKSHOP AGENDA
LIST OF PARTICIPANTS
LIST OF LIVING RESOURCES
TASK FORCE MEMBERS
ADDENDUM TO THE BENTHOS TECHNICAL
WORK GROUP REPORT
ADDENDUM TO THE SHELLFISH
PLANNING SESSION AND TECHNICAL
WORK GROUP REPORT
GENERAL COMMENTS ON THE MATRIX
APPROACH TO DEFINING HABITAT
REQUIREMENTS
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PREFACE
Finfish, shellfish, waterfowl and submerged aquatic
vegetation have declined in the Chesapeake Bay. Initial Bay
restoration efforts have focussed on improving water quality.
However, there has been growing recognition that the living
resources themselves may be the best guides to developing a
strategy for their recovery.
In 1986, the Chesapeake Bay Implementation Committee
established a Living Resources Task Force of managers and
scientists from Federal and State regulatory and resource
agencies, private industry and universities. The Task Force
was charged with the goal of developing a resource-based
approach to defining water quality and habitat objectives for
restoring and protecting living resources in the Bay. These
objectives would provide a framework for priority planning and
development during and following Phase II of the Chesapeake Bay
Program.
Through a series of meetings, the Task Force members
developed the following approach to setting resource objectives:
• They identified key representative species in Chesapeake
Bay. (Species were selected based on commercial and
recreational importance, declining populations and/or
importance to the Bay ecosystem.)
• They established priorities for immediate action among
these species.
• They identified the critical life stage of each Priority
I species within the Bay (i.e., the portion of the
species' life cycle thought to be most susceptible to
environmental conditions in Chesapeake Bay habitats and
the stage that would most benefit from restoration
efforts).
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• They developed, in matrix form, habitat requirements for
critical life stages of Priority I species. The
matrices included environmental and anthropogenic
factors (e.g., salinity, temperature, toxics
concentrations) affecting the key species as well as the
major subordinate species required for food or cover.
The matrices were combined into a document called "Strawman
II: Living Resources Habitat Requirements for Chesapeake
Bay." Where possible, the matrices included specific criteria
thought to be protective of the key and/or subordinate
species. Although not themselves enforceable, these criteria
could be used to provide guidance in setting regulatory water
quality standards.
Approximately 60 scientists reviewed the Strawman II
document at a one-day workshop on February 24, 1987, in
Annapolis, Maryland. This report presents the results of that
workshop. In the morning, the participants divided into six
planning sessions: Benthos, Plankton, Submerged Aquatic
Vegetation, Shellfish, Finfish and Waterfowl/Birds. Following
these sessions, participants split into four technical work
groups: Finfish, Molluscan Shellfish, Crabs and
Waterfowl/Birds.
In the planning sessions, participants discussed the
general habitat requirements for species within the associated
trophic level. In the technical work groups, participants
reviewed the habitat matrices and developed recommendations for
enhancing and refining the matrices. At the end of the
workshop, the chairmen gave brief reports on the achievements
and recommendations of their groups.
This report, divided into seven sections, summarizes the
proceedings of the workshop. Each section presents the
recommendations and conclusions of a planning session and/or
work group (sessions and groups that dealt with the same
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species have been combined). The workshop agenda, list of
participants and a list of Living Resources Task Force members
are presented in Appendices A, B and C respectively. Appen-
dix D presents the revised habitat requirement matrices for
target species and supporting trophic food species. Appendix E
is an addendum to the report of the benthos planning session.
Finally, Appendix F lists general comments on the habitat
requirements matrices approach.
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1. BENTHOS PLANNING SESSION
The Benthos Planning Session was chaired by Dr. Fred
Holland, of Versar ESM Operations. The conclusions of the
group are presented in this section. Following the workshop/
Dr. Holland submitted an addendum, included as Appendix E, that
provides more detail on the planning session report.
Benthic communities are an integral part of the food web of
Chesapeake Bay and serve an important role as habitat formers.
Benthic organisms actively change the nature of the Bay habitat
through such processes as bioturbation, nutrient
remineralization and structural modification. They directly
affect water quality through interaction with sediment and
water. That direct interaction makes benthic communities more
sensitive indicators and integrators of overall water quality
(particularly dissolved oxygen levels) than direct measurement
of water quality. They can also indicate relative sediment
quality, and are easily collected and enumerated.
Much of the upper Bay benthos (especially deeper portions)
is stressed, and is characterized by shallow burrowing, high
productivity and rapid turnover. The upper Bay benthic
communities have changed from filter feeding to predominantly
deposit feeding.
Fish and other predators affect the recruitment of benthic
organisms. The upper Bay benthic communities tend to consist
of small, fast-growing organisms with high turnover rates.
These species may not be the preferred prey for fish and
waterfowl (Holland, 1986). Abundances of estuarine benthic
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species have been increasing since 1970 (though this may not be
true for the benthic community in deeper waters) (Holland et
al., 1984).
Habitat criteria can be defined for the benthic community;
however, synergism among the parameters must be considered.
For example, tolerance to salinity may change as temperature
changes-. At the extremes of an acceptable range, organisms
become very intolerant. The matrix approach in the Strawman II
document does not consider potential synergism between various
habitat parameters.
The group pointed out that it is important to define
management goals for benthos. Managing for benthic production
would imply a eutrophic (but not polluted) system, whereas
managing for a variety of species would require a different
approach.
Session participants felt that it would be easy to identify
the groups of benthic species that are representative of
various specific habitats along the Bay. However, they had
difficulty with the concept of establishing water quality
parameters based on a critical life stage. The group felt that
it was more appropriate to manage for population success as a
whole than for the the success of any individual part of the
population. Participants pointed out that there may be
different critical life stages in different regions of the Bay
for the same organisms.
In addition, some participants noted that fish eat many
different kinds of benthic organisms. Thus, it makes more
sense to identify water quality parameters that will protect
benthic organisms as a whole than to do this on an individual
species basis.
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Following the workshop, Dr. Holland submitted an additional
paragraph for the Benthos Planning Session Report:
Over the last several decades the character of Chesapeake
Bay benthic communities has changed. Filter-feeding
benthic organisms, including oysters and shoft-shelled
clams, have generally become less abundant, and small,
rapidly growing deposit-feeding species have become more
abundant. Recent increases in the abundance of
deposit-feeding benthos appear to be associated with
long-term changes in Bay water quality, especially
increased nutrient levels and algal productivity (Holland
et al. 1984). As algal productivity has increased so have
organic detritus inputs to bottom habitats. This detritus
is the preferred food for deposit-feeding benthos. Because
benthic organisms are important prey in the diets of
commercially and recreationally important fish and
waterfowl, recent changes in the character of benthic
communities may be one factor contributing to recent
declines in abundance of some fish species (e.g., white
perch and striped bass) and increases in abundances of
other (e.g., spot). Small, rapidly growing deposit feeders
are a suitable prey for spot but may not be as suitable for
striped bass or white perch.
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2. PLANKTON PLANNING SESSION
2.1 Introduction
The Plankton Planning Session was chaired by Dr. Kevin
Sellner, of the Benedict Estuarine Research Laboratory. The
participants produced the following report.
As the basis for further discussion, the group agreed that
the Chesapeake Bay is a plankton-based ecosystem. Therefore,
plankton, as the food source for production of critical life
stages of the "key species," control overall fish and shellfish
biomass in the Bay. The Strawman II document considered
plankton primarily as supporting food chain habitat components
of "key species." Session participants recommended that the
process for developing habitat requirements emphasize that the
Bay is a trophic system where all organisms are inextricably
linked to the plankton. The group suggested that plankton
control of "key species" production implies that plankton are
the key organisms in the system. Factors that control
fluctuations in plankton numbers, sizes and production
(including circulation patterns in the Bay and tributaries) are
critical to the success or failure of "key species."
Therefore, the Task Force should consider "key species"
production from the lowest trophic levels up, rather than from
the top predators down.
In this endeavor, the Plankton Planning Session offered
four hypotheses for control of "key species" production to be
considered by the Living Resources Task Force.
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2.2 Hypotheses
a. Metazoan Food Web
Production of "key species" in the Strawman II document
considers classical food web theory, i.e., that fish and
shellfish production is a result of a metazoan food web
(simplistically, carbon transfer from phytoplankton to copepods
to fish). The planktonic food web of Chesapeake Bay includes a
microbial-based web as well as the classical metazoan food web
implicit in Strawman II food chain requirements. There is
growing evidence that a combination of factors - probably
arising from synergistic effects of point and nonpoint source
additions of nutrients (eutrophication) and toxics - may be
resulting in high bacterial production and an abundance of
small phytoplankton taxa. A well-developed microbial food web,
including high densities of small microzooplanktonic suspension
feeders, is associated with high oxygen demand, loss of aerobic
habitats and, possibly, an altered food web that would reduce
production in the highest trophic levels (key species).
b. Impact of Key Species on Plankton Dynamics
Several pelagic taxa overlooked in the Strawman II document
consume large quantities of plankton, leaving less planktonic
substrate for "key species" production. The impact of the bay
anchovy (the most numerous Bay fish) and ctenophores/jellyfish
on plankton dynamics should be considered in potential
production of the "key species" listed in the Strawman II
document. Bay anchovy affect the system because they may
consume large portions of the available plankton prey,
diverting much of the carbon away from the key species.
Ctenophores and jellyfish are major consumers of zooplankton
prey and larval fish in the system.
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c. Effect of Nutrient and Toxics Loadings
Chesapeake Bay plankton respond most rapidly to subtle
changes in nutrient and toxic loadings from anthropogenic or
environmental sources in the watershed. These changes may
include alterations in the size and species composition of
plankton communities from "normal" assemblages characteristic
of the system. Increased production of the
perturbation-selected taxa may divert carbon away from key
species by modifying classic trophic linkages, possibly
contributing to lower production of "key species." Thus, it is
important to focus management decisions on the control of
anthropogenic inputs that will alter normal "suites" of
plankton.
d. Correlation of Larval Stages with Plankton Density
Maximum survival of larval stages of "key species" should
be correlated with highest densities of microzooplankton (20 to
200 micrometers) and mesozooplankton (greater than 200
micrometers) in the Bay.
2.3 Conclusion
Chesapeake Bay is a phytoplankton-based system. Any
initiatives favoring selective growth of a "healthy"
phytoplankton assemblage will maximize potential production of
desirable living resources (key species) in the Bay.
Data are needed on the carbon demand for critical life
stages of key species in order to estimate whether plankton
populations are adequate to support these species. Data are
also needed on the selectivity of key species at their critical
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life stages: Do they prefer certain size or species of
plankton? Do they require a specific food quality for proper
development, e.g., high protein, high lipid, high
carbohydrate? Data are also needed on the temporal and spatial
distributions of all plankton types and the critical life
stages of the key species of the higher trophic levels.
The overall recommendation of the Plankton Planning Session
is that, since plankton are the "key" to Bay fish and shellfish
production, the Task Force should concentrate on environmental
and anthropogenic factors that control availability of plankton
for estimating the success of critical life stages of "key
species" in the Bay.
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3. SUBMERGED AQUATIC VEGETATION PLANNING SESSION
3.1 Introduction
The planning session was chaired by Dr. Court Stevenson,
Horn Point Environmental Laboratories. The group determined
that submerged aquatic vegetation (SAV) can be divided into
three groups that have different water quality requirements:
plants in high salinity areas (e.g., Zostera marina and Ruppia
maritima), plants in mid-salinity (Potamogeton pectinatus,
Potamogeton perfoliatus, Ruppia maritima and Zannichellia
pulustris), and plants in low salinity to tidal freshwater
(Hydrilla verticillata, Heteranthera dubia, Myriophyllum
spicatum, Ceratophyllum demersum, and Vallisneria americana).
3.1.1 High Salinity
The high salinity environments in the lower Bay tend to be
more nitrogen-limited. When nitrogen concentrations are high,
algal growth is a problem for SAV. Dense phytoplankton blooms
shade submerged aquatics as well as promote algal epiphytes
which can form dense colonies on the leaves. The current view
is that epiphytic and epifaunal overgrowth can weaken submerged
aquatic populations by limiting primary productivity through
shading, thus depleting carbohydrate reserves. If substantial
carbohydrate energy is not stored throughout the winter in
subsediment, roots and rhizomes, growth of SAV will be
adversely affected in the spring. If algal epiphytes continue
to overgrow the plants for several years, this can cause a
decline in SAV, as observed in the late 1970s.
Both nitrogen and phosphorus seem to stimulate SAV growth
in the high salinity region when applications are made in the
root zone.
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3.1.2 Mesohaline
The mesohaline environment has a gradient of nutrients,
with high levels at the heads of estuaries to relatively low
levels at the midpoint of the Bay. There has been a resurgence
of SAV growth in recent years, particularly in the mesohaline
areas with elevated salinity levels. Data indicate that SAV
populations may decline indirectly due to overenrichment at
average summer concentrations in the water column of greater
than 0.14 mg/1 dissolved inorganic nitrogen and greater than
0.01 mg/1 phosphate. Thus, levels of less than 0.14 mg/1
dissolved inorganic nitrogen and less than 0.01 mg/1 phosphate
may be suitable for use in defining areas that will support SAV
growth in brackish waters. These values appear to be
thresholds at which epiphytic overgrowth becomes problematic to
SAV. Light conditions in the mesohaline portions of the Bay
are often limiting, particularly in the summer. The group
recommended (1) that attenuation coefficients should not exceed
a Kd of 2 (photosynthetically active radiation - 400 to 700
nanometers); (2) that levels of suspended solids in the water
column levels should be less than 20 mg/1, and (3) that
chlorophyll a in the water column should be less than 15 ug/1.
3.1.3 Freshwater
Substantial regrowth of freshwater SAV has occurred over
the last several years in tidal fresh portions of the Potomac.
This is probably due in part to the reduced nutrient loading
from the Blue Plains wastewater treatment plant, which caused
decreased algal growth, hence less shading in the water column
and via epiphytes. Also, low runoff in 1985 and 1986 caused
decreased nonpoint-source nutrient loadings which appear to
have increased SAV growth. Participants felt that the current
regrowth was an excellent natural experiment that should be
analyzed further to provide data on the relationship of
nitrogen and phosphorus levels to SAV growth.
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There can also be substantial SAV growth at the head of the
Bay in high nitrogen concentrations (in the range of 0.7 to
1.4 mg/1) as long as phosphorus concentrations are very low
(less than 0.01 mg/1). SAV can grow in part because the low
phosphorus inhibits algal growth, and the SAV can obtain
phosphorus from sediments. However, some SAV species may
create high enough daytime pH levels to activate release of
phosphorus from sediments, thus causing algal blooms. This
mechanism may be partially responsible for the high pH in the
Potomac estuary.
3.2 General Comments/Recommendations
Light intensity in the Bay is less than it has been
historically, and it was felt that a return to pre-Agnes (i.e.,
prior to June 1972) levels was a worthwhile goal. One source
of information for these levels is Effects of Tropical Storm
Agnes on the Chesapeake Bay Estuarine System (Davis et al.,
1976).
Nutrients and sediments limit SAV more than low salinity.
Phosphorus is an important limiting factor, particularly for
epiflora in shallow freshwater. Nitrogen could also be an
important factor in higher salinity areas. It was pointed out
that nutrients must be considered together, and that nitrogen
as well as phosphorus should be considered in management
decisions on reducing nutrient inputs to the estuarine portions
of the Bay.
At present, data indicate that metal concentrations in
Chesapeake Bay sediments are not high enough to be toxic to
SAV. Submerged aquatics can sequester metals in their tissues
and serve as indicators for past pollution episodes.
Herbicides do affect SAV. Widely used herbicides such as
atrazine may have local effects on submerged aquatics in
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shallow embayments that are affected by agricultural runoff.
It was tentatively agreed that levels less than 10 ppb would
not present a problem in open waters. The journal literature
does have information on specific herbicide levels that impact
particular SAV species.
Pesticides do not appear to harm SAV directly, but they do
adversely affect invertebrates and heterotrophic food chains;
thus potentially harming SAV. For example, pesticides may
adversely impact snails (Bittium sp.), which usually clean
epiphytes from leaves. Declines in snail populations could
cause reduced photosynthesis for the plants.
The group agreed that transplantation of submerged aquatic
plants provides an excellent environmental measurement of
existing water quality. These transplantation efforts should
be closely monitored to elucidate the relationship between
water quality and continued reestablishment of SAV.
As much new literature on SAV has been published recently,
the group recommended that the comprehensive literature review
conducted by Dr. Stevenson (Stevenson and Confer, 1978) be
updated. The U.S. Fish and Wildlife Service is considering
funding this.
Participants recommended that increased emphasis be placed
on habitat monitoring of water quality, particularly in the
more shallow SAV beds. This monitoring would serve to document
continuing changes in water quality in an effort to define
population requirements in various sections of the Bay.
A report being prepared by Court Stevenson and Lorie Staver
for the Maryland Department of Natural Resources Tidewater
Administration will provide information on water quality
parameters associated with the resurgence of submerged aquatic
vegetation in the mid-Chesapeake. The report will be available
in July 1987.
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4. SHELLFISH PLANNING SESSION AND TECHNICAL WORK GROUP
4.1 General Approach and Recommendations
The planning session and technical work group were chaired
by Dr. Roger Newell, Horn Point Environmental Laboratories.
Reports from both groups are combined within this chapter.
Participants agreed that many estuarine species of bivalve
are similar in their tolerance of certain environmental
parameters, e.g., suspended solids, dissolved oxygen.
Therefore, the group developed comments and recommendations for
each parameter that would generally apply to all molluscan
shellfish. Separate criteria should be developed only when
there is a real difference in response between species, e.g.,
substrate type. The similarities between species mean that
creating conditions that are favorable to one species will
generally benefit other species.
Participants recommended that interactions between
parameters be considered. They cautioned that single factor
analysis would never be sufficient. For example, an animal
might be unaffected by one factor alone but synergism or the
additional sublethal stress provided by a second may result in
a reduction of fecundity or larval viability.
The group commented that it was unrealistic to try to
restore the Bay to its former condition, i.e., that existing
prior to colonization by European settlers. Instead, emphasis
should be placed on resource management to try to retard the
accelerated pace of change to the system and explore
enhancements of fishery habitats in more localized areas.
Further discussion and consideration are necessary to establish
a desirable and realistic goal for mollusc population size.
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4.2 Geographic Distribution
Participants recommended that management goals should aim
to expand the range of all species up to their tolerance
limits, especially into low salinity regions. This would
require limiting harvest pressure in the low salinity areas.
Where possible, sanctuaries should be maintained in marginal
habitats. This might help to provide a reserve of individuals
that would be available to colonize the more optimum habitats.
The optimum habitats should also be preserved and managed to
help modify the effects of fishing pressures.
The group considered the importance of diseases (e.g.,
Haplosporidum nelsoni [MSX] and Perkinsus marinus [dermo]) and
predators in controlling the oyster population and
distribution. Although these factors cannot be controlled at
present, they do regulate geographic distribution of species.
Natural factors, including diseases, predators and climatic
variation, have a much greater influence on oyster populations
than anthropogenic and environmental factors that can be
controlled by management practices. This should be taken into
account when making management decisions.
Commercial harvesting has changed the oysters' habitats.
Dredging and overharvesting have spread out or reduced the
height of the reefs. The reefs are now broader and have much
less relief above the sediment and are, therefore, more
susceptible to sedimentation processes.
4.3 Critical Life Stage and Period
The group agreed that both the larval and adult life stages
for clams and -oysters are critical life stages, and that each
stage is susceptible to different stress factors.
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4.4 Habitat Requirements
Food (Including Chlorophyll, Nitrogen, N/P Ratios and
Carbon)
The Strawman II document gave different food requirements
for different species. Participants felt that this could be
simplified since all bivalves have very similar food
requirements. The group noted, however, that there is a
critical food size for different life stages. (The group
discussed the importance of involving phytoplankton experts in
developing a strategy to manage the environment so as to
maximize the production of 3- to 35-micrometer (diameter) cells
that bivalves feed on. The group suggested that it was
important to understand how any changes in the patterns of
primary production that may be occurring in Chesapeake Bay
affect all life stages of the molluscs (see Section 2.1).
Rather than separately consider major food species,
chlorophyll, nitrogen, N/P ratios and carbon, participants
considered them together as a single food requirement. The
group recognized the complexity of factors affecting primary
production and concern with the statement made by the Plankton
Planning Session (see Section 2.1). They cautioned that, for
some criteria, size must be considered to ensure availability
to the animal. For example, chlorophyll in cells smaller than
3 micrometers will not be available to the animal; thus a total
chlorophyll measurement could be deceptive. Chlorophyll
measurements should, therefore, be partitioned into the
appropriately sized fractions.
Substrate, Suspended Solids, Turbidity, Secchi Depth and
Light Intensity
The group agreed that sedimentation (including substrate,
suspended solids, turbidity, secchi depth, and light intensity)
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is an important habitat factor influencing the continued
propagation of shellfish. Members felt that the principal
factor affecting the success of the oyster in Chesapeake Bay is
the lack of clean cultch. Clean oyster shells (cultch) are the
preferred substrate for oysters, and oyster larvae require a
clean shell for settlement and metamorphosis. However, factors
such as heavy harvesting and disease are causing a decline in
the preferred substrate. An active oyster bar is not subject
to siltation because it extends into the water column where
currents carry away biodeposits and silt. However/ once the
bar has been compromised (e.g., by overharvesting and high
sedimentation rates), the system shifts from a filter-feeding
system to a deposit-feeding system. It then becomes very
difficult to return the bed to its former condition. Some
participants questioned whether changes in sedimentation rates
would have much effect on deposit-feeding systems. The group
discussed using sediment trap methodologies to estimate
sedimentation in the actual oyster-producing areas. These
methods would provide better measures of water quality than
turbidity or secchi depth.
Participants stressed that it was important to manage the
Bay to reduce loads of suspended particulate inorganic
material, especially during the period of spawning and larval
settlement. Repropagation of SAV beds in critical habitats
would reduce resuspension of bottom sediments but would not
prevent deposition through the water column. Adult mobile
infaunal clams are not as sensitive to burial by sediments as
oysters, but juvenile clams can actually be smothered by
siltation.
Participants commented that the Bay is a pulse system with
many fluctuations (see Appendix E), and therefore it would be
difficult to set a specific level for total suspended solids.
The group criticized the sediment-related criterion of
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1,000 mg/1 as being unrealistic, probably because it was
derived from dredge areas rather than the natural environment.
Cover
SAV cover is important in reducing turbidity within the
system, thereby maintaining cultch quality. SAV also provides
a very important refuge for juvenile clams from crab and fish
predation. Reestablishment of SAV could markedly increase
production of infaunal clams. Oysters are the most important
cover for setting oysters.
Temperature and Salinity
Participants did not attempt to set levels for these
parameters. Temperature and salinity fluctuations are normal
habitat requirements. Anthropogenic effects on Bay conditions
are not significant enough to make these parameters worth
considering as a management issue except in local areas, e.g.,
power plant discharges, which are already strongly regulated.
Anthropogenically altered freshwater flow to the estuary might
modify both temperature and salinity and should be considered.
Metabolic activity of shellfish is strongly temperature-
linked and must be considered in relation to other parameters.
Environmental factors are less important in the winter when the
shellfish are dormant. Participants noted that larval stages
are more sensitive to temperatures.
Flow
Brief salinity fluctuations that result from natural flow
patterns may aid the control of parasites and disease (see
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Appendix E). Therefore, it might be beneficial to reestablish
oyster bars in areas where flow patterns would encourage
periods of low salinity.
£H
The group changed this parameter to "6.8 to 8.5," but noted
that pH fluctuation is a natural phenomenon and would be
difficult to control. pH is lower under anoxic conditions;
therefore, steps should be taken to control anoxia. Changes in
pH may affect the phytoplankton community, which in turn will
affect molluscs.
Dissolved Oxygen
Dissolved oxygen (DO) is critical for all molluscan
shellfish life stages. However, the tolerance of anoxia varies
with life stage and with season. In the summer, the tolerance
is markedly reduced. Participants recommended that the habitat
matrices have a seasonality component, and that a matrix of
these interactions be developed. Data are needed on how long
species can survive under anoxic conditions. Ongoing research
as part of the NOAA/Seagrant Hypoxia Program will provide new
insights, particularly concerning the effect of low DO on
larvae. Lateral movements of anoxic bottom waters over clam
and oyster beds and the impacts on these beds should be studied.
Ionic Constituent
The group determined that this parameter is not applicable
to molluscan'shellfish.
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Bacteria
Participants recommended that this parameter be retitled
"Pathogens." They discussed the importance of pathogens in
controlling geographic distribution (see Geographic
Distribution above).
Phosphorus
This parameter is not directly applicable to molluscan
shellfish, but could become a factor through the food chain,
PAHs, Metalsf Insecticides, Herbicides and Chlorinated
Hydrocarbons
The group considered these classes of compounds together.
Tributyltin was also mentioned by one participant as being a
toxicant of concern. Although there are good data to show that
all these compounds can be highly toxic, especially to larval
stages, the general consensus was that these compounds may not
be that important in regulating production on a Bay-wide
basis. The group felt that current efforts to enforce existing
toxicant standards should be adequate for protecting oyster and
shellfish populations. It is possible, however, that toxicants
may pose a problem in local environments where toxicants are
discharged into the Bay (e.g., localized use of antifouling
compounds). In general, however, toxicants sequestered in
shellfish tissue are a human health concern, if consumed,
rather than an important influence on shellfish survival.
Participants thought that the metals levels listed in the
Strawman II document would protect larvae, and that small
molluscs may tolerate even higher levels. The group questioned
the criterion of less than 0.0001 ppb for mirex. They asked
-18-
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that this figure be double-checked. Oil in the Sea/ Inputs/
Fates and Effects (National Research Council, 1985) was cited
as a reference for effects of hydrocarbon. Another reference
that may be of interest is an NAS report on detergents used to
clean up oil spills/ to be published in 1987.
Following the workshop, Dr. John Kraeuter (Baltimore Gas
and Electric Company) submitted the following statement:
While specific effects of oil on oysters/ hard clams and
soft clams have been shown, these data are derived mostly
from information collected in conjunction with major oil
spills. The effects of oil at low concentrations are not
as well known, but developmental processes can be sensitive
to petroleum/ and even fairly low concentrations can result
in measurable abnormalities (less than 1 mg/1). Hydro-
carbons also have histopathological and/or mutagenic
potential, and concentrations of petroleum as low as 10
ug/1 can alter normal behavior of many marine organisms.
In view of the National Academy Review, the Shellfish group
recommends efforts to reduce petroleum hydrocarbon input to
Chesapeake Bay. It would seem this can best be done by
controlling outputs in ports, marinas and harbors (boats),
runoff from storm drains (streets and roads) and from
municipal wastewater facilities.
Hardness and Alkalinity
The group agreed that these parameters are not applicable
to molluscan shellfish.
Other Comments
Following the workshop, Dr. Kraeuter submitted a statement
on the importance of Bay processes. This is included as
Appendix E.
-19-
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5. FINFISH PLANNING SESSION AND WORK GROUP
This report covers the finfish planning session and
technical work group/ which considered both anadromous and
marine spawning finfish. The sessions were chaired by Dr.
George Krantz, Maryland Department of Natural Resources.
5.1 Species List
The planning session began by discussing changes to the
list of priority finfish species in the Strawman II document.
The changes that participants made to the list are indicated in
Table 1. The group thought that the Strawman document should
include any ecologically important fish, regardless of their
current level of prosperity or commercial or recreational
significance. The bay anchovy and killifish were added to the
Priority I list because of their ecological importance. The
killifish is extremely important for nutrient exchange between
the marsh and the higher fish food chain. The hog choaker was
added because it is the most pollution-tolerant species and can
therefore act as an indicator of degraded environments. Any
damage to this species would suggest that the more sensitive
species are seriously threatened. (The hog choaker would
therefore not be appropriate to use as a basis for modelling or
management.) The Atlantic sturgeon was moved from Priority I
to II because there are so few of them that participants did
not think it appropriate to base management decisions on this
species. The croaker was moved from Priority I to II because
its population fluctuations are not thought to be directly
related to the Bay. The naked goby and oyster toadfish were
added to Priority III because they are an important ecological
link. Cobia was dropped from the Priority II list because no
one in the work group could attest to the importance of this
ocean fish to the Bay.
-20-
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-21-
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5.2 General Changes and Recommendations
Participants agreed that the background information
supplied for each species was unacceptable. They recommended
that new background writeups be prepared based on A.J.
Lippson's Bay Atlas (see Table 2 for reference). They also
recommended this reference as a good source of information on
geographic distribution. Several other sources were identified
which contain species distribution and spawning ground maps,
including Habitat Sensitivity maps for Maryland, Corps of
Engineers Map Folio, etc. (see Table 2). The group agreed that
the terms, categories and citations in the matrix should be
clarified.
The "critical life stage" was defined as the period in
which habitat variation has the greatest impact on a given
species. For each species discussed, the group reviewed the
criteria and indicated whether they were critical (i.e.,
essential to survival), noncritical, or tentatively critical
(not of concern at current levels, but potentially critical to
survival if present environmental conditions are altered).
Participants asked that it be noted in the Strawman document
that all the listed habitat criteria have some important
biological impact at some level, even though this level may
appear extreme compared to present ambient levels. They also
noted that negative synergistic effects could become evident at
the upper and lower limits for any parameter. Synergistic
effects, though not considered in the current matrix approach,
could radically alter any fish species response to a specific
habitat criterion. For example, hardness by itself is not
considered critical, but in combination with low pH and heavy
metals, the synergistic effect is fatal. The finfish group
felt that synergistic and interaction effects would become more
critical when habitat conditions approach the margin of
tolerance of any parameter.
-22-
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TABLE 2
INFORMATION SOURCES FOR FINFISH
1. U.S. Fish and Wildlife Service - Species documents
providing habitat suitability curves for individual
species (i.e., shad, striped bass) to be used in IFIM.
2. Habitat Suitability Index Documents - Biology Report.
National Wetlands Center (formerly National Coastal
Ecosystems Team), U.S. Fish and Wildlife Service, Slidell,
Louisiana.
3. Atlantic States Marine Fisheries Commission - Management
plans for a number of species (those listed as Priority I)
4. U.S. Corps of Engineers (COE) - New England region
species-specific biology profiles.
5. Susguehanna River Anadromous Fish Restoration Committee -
Restoration of American Shad to the Susquehanna River,
1986 Annual Progress Report. U.S. Fish and Wildlife
Service, Harrisburg, Pennsylvania. 340 pp.
6. U.S. Army Corps of Engineers (COE), 1982 - Map Folio:
Chesapeake Bay Low Freshwater Inflow Study, Phase II,
Biota Assessment. Prepared for the U.S. Army Engineer
District Baltimore, by Western Eco-Systems Technology,
Inc. 204 215th Street, Bothell, Washington 98011.
7. Virginia Institute of Marine Science Anadromous Fish
Project Annual Reports.
8. National Oceanic and Atmospheric Administration (NOAA)
Sensitivity Maps: Ann Hayward-Walker of NOAA was
mentioned as someone who is updating various species
sensitivity maps and atlas information and whose work
might be included in the final document.
9. University of Maryland - Baird/Ulanowicz (authors),
"Chesapeake Ecosystem Network Documentation."
10. New Orleans Coastal Ecosystem Studies.
11. Ronald Hellenthal's Trophic Information (University of
Notre Dame, Department of Biology, South Bend, Indiana
46556).
-23-
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TABLE 2 (continued)
12. Draft Report on Polynuclear Aromatic Hydrocarbons and the
Chesapeake Bay, Maryland Department of Natural Resources,
October, 1986.
13. West in, D. and B. Rogers (1978) - Synopsis of Biological
Data on the Striped Bass, Morone saxatilis (Walbaum)
1792. Univ. of Rhode Island Marine Tech. Report 67.
14. A.J. Lippson (1973) - The Chesapeake Bay in Maryland: An
Atlas of Natural Resources. The Johns Hopkins University
Press, Baltimore and London. 55 pp.
-24-
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The group worked thrown ' nc nu-t/i
alewife/blueback herrinq, i.iu .1 :.anti<.
established criteria for n._ > •• i i i.-al
species. Participants dj-,; L..;. t,y tt,-
range for the Priority j c( \.-v- ;.-.," ..iil
Rather, members tried t.o csr. <-, u i > ;;h a <.
ces for the striped bass,
menhaden and spot, and
Jife stage of these
establish an optimum
icdi life stages.
<.,iiterion level or range
that would ensure no adv wiictt the fish need, not what
habitat conditiont> they -~u; : ^u' iy .,,ni \-ive in. Thus, in some
cases, current data inay uc. ; ua^ui \.jji i at t.
Participant, s si roiiv) s ;
might not be relevant i >.,
deleted from the stra^iad*
preamble state that the
protecting species; i.e.
exceed the criteria, Pdi
the potential for misuse
taken as allowable leveJ
protect species.
^i .,1-1 (i<. ,i ji^en represent the extremes for
luthsid! conditions should never
;oii-'an!b were very concerned about
r ^iileria; i.e., criteria could be
for degradation rather than limits to
Much information in the lileiature was not in the matrix.
Since no single definitive coun'e document exists, the group
recommended that several additional documents be attached to
the Strawman document to pri'^jde background information on the
elements of the matrix (:-ee Table 2),
After working through the f ? , st two species, participants
found that some parameters were generic to specific groups of
finfish in similar trophic levels,, They discovered that all
the matrix elements influence trophic dynamics, and therefore
supported the conclusions of the plankton work group.
Conceptually, management of the habitat for fishes must include
all parameters that, sustain intermediate trophic levels and
near ideal conditions.
-------
Within the matrices, several changes were suggested. The
"Bacteria" criterion should be changed to "Pathogens." The
"Zone" criterion should be specified as "Vertical Zone," since
horizontal zone is covered by geographic distribution. The
"N/P Ratios" criterion should be placed next to the "Nitrogen"
and "Phosphorus" parameters in the matrices. "Chlorine" should
be added as a critical variable to all matrices. The group
suggested that, if the tables are to be used for management
decision-making, they should exclude all factors that are
uncontrollable by existing management techniques, although a
number of parameters (i.e., flow, temperature) can be critical
to survival and can be affected by development, dams, industry,
etc.
Several habitat parameters can be treated generically.
Biological systems in fish have similar basic requirements and,
therefore, similar responses to most environmental features.
Participants did not set any levels for nitrogen and phosphorus
for any species, but requested that the following points be
noted concerning these parameters: Ammonia, nitrites and any
form of reduced nitrogen are known to be toxic. Nitrogen and
phosphorus can have direct toxic effects on finfish, but the
most critical impact is their collective effect on food
production and anoxia in stratified waters. These factors must
be taken into account when setting levels for these parameters
for finfish.
Temperature, pH and dissolved oxygen could be treated
generically with a few minor exceptions by species or by
geography of species. Habitat levels of 6.5 to 8.5 for pH and
greater than 5 mg/1 for dissolved oxygen were felt to be
acceptable as generic criteria.
Metals, PAHs, chlorinated hydrocarbons, herbicides and
pesticides were combined for all Priority I species. The group
-26-
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decided that levels that would have no adverse impact on the
most sensitive species in this group (the alosid) should be
used for all species in the absence of species-specific data.
Again, the philosophy was stressed that levels should be set so
that there is rio biological impact. The group stated that the
levels set should minimize the possibility of intertrophic
magnification or additivity of toxic chemicals as a result of
chronic exposure. However, the bay anchovy and killifish are
more tolerant so criteria could be higher for these species.
The Interstate Fisheries Management Plans of the Atlantic
States Marine Fisheries Commission (ASFMC) were cited as a
source for data on these parameters (ASMFC, 1985; Atlantic
Menhaden Management Board, 1986). Data from the Draft Final
Report on Polynuclear Aromatic Hydrocarbons and the Chesapeake
Bay (Maryland Department of Natural Resources, 1986) suggest
that fish experience lethal effects when exposed to 12 ppb PAHs.
Following the workshop, Dr. Krantz submitted the following
statement regarding the matrix approach:
Relationship between fish habitat and their population
success must follow closely the concept of the weakest link
in the chain of life. The matrix exercise has focused on
critical criteria and may be missing the concept that if
any single habitat criterion is violated, the chain, with
all its intact critical links, would still be broken.
Though often difficult to comprehend, this concept is
essential and must be considered in planning. Two very
important axioms must be considered.
First, one adverse experience in the life cycle of a fish
population can be critical. For example, transit
phenomena, which are of very short duration (minutes), can
destroy the chain at any point over the course of the
entire life cycle of a fish species. For mathematicians,
this means that averages cannot be used, only the extreme
of the numerical distribution. This is the reason the
group suggested upper and lower incipient levels for
responses.
The second axiom is that habitat parameters for finfish
must also include the most constraining value for every
other component of the trophic ecosystem. The critical
chain of life with its weak links also runs vertically
-27-
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through the trophic levels that are used to describe
biomass or energetics (nitrogen, phosphorus, and carbon).
These components must be passed successfully through the
pyramid to the higher trophic level occupied by fish. A
diminution of a lower trophic level (algae, zooplankton,
worms) would have an adverse impact on the higher trophic
level. What escapes many scientists and the lay manager or
planner is that a small change in a lower trophic level is
multiplied by each trophic level that a contributing
trophic component is passed through. For example, a very
small change (e.g., a 10 percent change in this example) in
an algae species that is consumed by a food chain that
reaches the fifth trophic level of a given fish species is
technically raised a minimum of five times when its impact
is ultimately expressed in fish biomass. This means that
we could expect a potentially large change (e.g., a 50
percent change in this example) in fish biomass from a 10
percent change in an important algae trophic component.
Unfortunately, the human element focuses on commercially or
recreationally important species. We have failed to
realize that insults to the lower trophic level organisms
are magnified by passage through the food chain. This
phenomenon alone could explain the demise of many of the
Bay's fish species. Therefore, all habitat criteria that
have a detrimental effect on any trophic level should be
described as negative factors in this matrix exercise.
These negative factors may not fit the matrix format now
being used for fish if they occur in another trophic
level. If a lay manager focusses only on fish, he or she
will fail to detect critical criteria in other trophic
matrices.
5.3 Matrices
Matrices were filled out for three species: striped bass,
alewife/blue herring and Atlantic menhaden. The above
consensus point, that information on the alosids provides
protection to all species, evolved by comparing the results of
these three species to all others.
5.3.1 Striped Bass
Background. Dr. Krantz recommended the Guidelines for
Striped Bass Culture (Bonn et al., 1976) as a source of
-28-
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background information. In addition, the group compiled a list
of nine references on the striped bass (see Table 3). Grant
and Olney (1982), Grant and Olney (1981) and Olney et al.
(1983) provide patterns of abundance of eggs and larvae in the
James, Pamunkey, Mattaponi, and Rappahannock rivers. Tresselt
(1952), Massmann et al. (1952, 1962), Rinaldo (1971) and
Merriner et al. (1980) provide additional documentation of
spawning activity.
Critical Life Stage. The critical life stage was expanded
to include both larval and juvenile stages.
Critical Life Period. The critical life period for the
striped bass was discussed. The group agreed that the critical
life period for larval and juvenile stages is April to June.
Food. Prey was not discussed for any species.
Substrate and Cover. The group decided these parameters
were not applicable to the critical life stages of the striped
bass.
Zone. Zone was changed to "water column, demersal."
Salinity. Salinity was reduced to "0 to 5 ppt" for the
critical life stages.
Flow. The group agreed that flow is a critical parameter
and reduced it to "0 to 0.5 m/sec." Flow velocity keeps
striped bass eggs and larvae suspended in the water column
which is their natural habitat. Lower flows would transport
the critical life stage out of the microenvironment needed for
proper development.
-29-
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TABLE 3. REFERENCES FOR STRIPED BASS
1. Grant, G.C. and J.E. Olney. 1981. Assessment of larval
striped bass, Morone saxatilis (Walbaum), stocks in Maryland
and Virginia waters. Part II. Assessment of spawning
activity in major Virginia rivers. Final Report, Segment 1,
to the National Marine Fisheries Service, Gloucester, Mass.
(Grant No. NA80FAD-VA1B), 39 pp.
2. Grant, G.C. and J.E. Olney. 1982. Assessment of larval
striped bass, Morone saxatilis (Walbaum), stocks in Maryland
and Virginia waters. Part II. Assessment of Spawning
Activity in Major Virginia Rivers. Final Report, Segment 2,
to the National Marine Fisheries Service, Gloucester, Mass.
(Grant No. NA81FAD-VA3B), 42 pp.
3. Massmann, W.H., B.C. Ladd and H.N. McCutcheon. 1952. A
biological survey of the Rappahannock River, Virginia. Part
1. Virginia Fisheries Lab, Gloucester Point, Virginia. 112
pp. (Mimeo).
4. Massmann, W.H., E.B. Joseph and J.J. Norcross. 1962. Fishes
and fish larvae collected from Atlantic plankton cruises of
R/V Pathfinder, March 1961-March 1962. Virginia Inst. of
Mar. Sci. Spec. Sci. Rept. No. 33, 20 pp.
5. Merriner, J.V., A.D. Estes, and R.K. Diaz. 1980.
Ichthyoplankton Entrainment Studies at Vepco Nuclear Power
Station. Final Technical Report 1975-19787. Va. Inst. Mar.
Sci., Gloucester Pt., Virginia. Section Ila and lib, 602 pp.
6. Olney, J.E., B.H. Comyns and G.C. Grant. 1983. Assessment
of larval striped bass, Morone saxatilis (Walbaum) stocks in
Maryland and Virginia waters. Part II. Assessment of
spawning activity in major Virginia rivers. Final Report,
Segment 3, to the National Marine Fisheries Service,
Gloucester, Massachusetts. (Grant No. NA81FAD-VA55B), 38
pp., Appendix I.
7. Rinaldo, R.G. 1971. Analysis of Morone saxatilis and Morone
americanus spawning and nursery area in the York-Pamunkey
River, Virginia. M.A. thesis, College of William and Mary,
Williamsburg, Virginia. 56 pp.
8. Tresselt, E.F. 1952. Spawning grounds of the striped bass
or rock, Roccus saxatilis (Walbaum), in Virginia. Bull.
Bingham Oceanogr. Coll. 14(1):98-110.
9. Maryland Department of Natural Resources. 1986. 1985
Striped Bass Status Report.
0993D
-30-
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Temperature. Participants agreed temperature is a critical
parameter. They noted that the criteria provided in the
Strawman II document were too extreme. At 12° C, the larvae
would stop growing; temperatures as high as 23° C would kill
them. The group changed this criterion to "16 to 19° C."
pH. This parameter was determined to be critical, but the
group agreed that not enough was known to set a criterion.
They pointed out that the level in the Strawman II document was
incorrect (pH = 6.5 causes great losses at low levels of
alkalinity). This is a prime example of synergism; research
has only recently detected this phenomenon.
Dissolved Oxygen. Participants agreed that DO is critical,
but not enough is known to set a minimal level with
confidence. A level of 5 mg/1 is known to have no adverse
effect on any life stage. Therefore, this level should be used
until additional research findings can further refine the
minimal level.
Ionic Constituent. The group was not sure whether ionic
constituent was a critical habitat criterion by itself. The
specific level in the Strawman II document was not discussed.
Turbidity and Suspended Solids. Turbidity was determined
not to be generally critical. Levels for turbidity and
suspended solids have been found not to be closely related,
even though these variables are normally linked.
Bacteria. Participants agreed that the category of
"Bacteria" should be changed to "Pathogens," since bacteria can
be an indirect food source. No one knew of any data suggesting
that striped bass larvae eat bacteria, so the group decided not
to include bacteria as a food source.
-31-
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Secchi Depth, Suspended Soli_dsL Light Intensity and
Chlorophyll. These parameters were determined not to be
critical. (See Turjbi^it£ above for note on suspended solids
level.)
Nitrogen and Phosphorus. Participants did not set any
levels, but requested that the following points be noted
concerning these parameters. "Ammonia, nitrites and any form
of reduced nitrogen are known to be toxic. Nitrogen and
phosphorus can have direct toxic effects on finfish, but the
most critical impact is their collective effect on food
production and anoxia in stratified waters. These factors must
be taken into account when setting levels for these parameters
for finfish."
PAHs. This parameter was de.=H gnated as a "provisional
critical parameter" pending more data. The group recommended
Westin and Rogers (1978) (see Table 2) as a potential source of
data on PAHs. Dr. Krantz also recommended reports from the
U.S. Fish and Wildlife Service (USFWS) Laboratory in Columbia,
Missouri, on toxicity tests on striped bass (Mehrle et al., in
press; Mehrle and Ludke, 1984),
N/P Ratios and Carbon. N/P ratios and carbon were
determined not to be critical to striped bass, but were primary
driving factors in trophic dynamics upon, which striped bass
depend.
Metals. This parameter was designated as a "provisional
critical parameter" pending more data. The group again
recommended Westin and Rogers (.1978) (see Table 2) as a
potential source of data on metals. Reports from the USFWS
Laboratory in Columbia, Missouri (Buckler et al., in press;
Mehrle et al., in press; Mehrle and Ludke, 1984)) were also
recommended. Aluminum and tributyltln were added to the list
-------
of metals of concern. Dissolved aluminum can impair gill
structure and efficiency in young striped bass. Low pH can
mobilize some metals. This is an excellent example of
synergistic effects that were not included in the matrix.
Hardness. This parameter was determined not to be critical,
Alkalinity. Participants agreed this parameter is critical
since it provides a buffering component to the ecosystem. They
changed the level given in the Strawman II document to read
"greater than or equal to 20 mg/1." They also noted that the
optimum range was 70 to 200 mg/1 calcium carbonate.
Herbicides/ Insecticides and Chlorinated Hydrocarbons. The
group decided that levels that would protect the most sensitive
finfish species (probably the alosid) should be used for all
species in the absence of species-specific data. Where data
allow, levels should be set so that there is no biological
impact. The levels set should minimize the possibility of
intertrophic magnification and additivity of toxic chemicals as
a result of chronic exposure. The chairman noted that, to
date, not a single compound could be identified as a problem in
the striped bass crisis. Dr. Richkus (Martin Marietta
Environmental Systems) supplied data on 96-hr TL "s for
striped bass larvae and juvenile striped bass for many
toxicants (Setzler et al., 1980).
Chlorine. Chlorine was added as a critical parameter, but
no levels were set (i.e., any amount is considered to be
detrimental).
Geographic Distribution. The group agreed that the
geographic distribution of striped bass should be "as Maryland
and Virginia have defined their spawning grounds by
regulation." This distribution would be less restrictive than
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the maps provided. Dr. Krantz will supply maps of the striped
bass spawning ground in Maryland. Dr. Barth (Virginia Marine
Resources Commission) supplied the Virginia Marine Resources
Commission regulation 450-01-0034 pertaining to the "taking of
striped bass." The description of the critical reaches is
provided in paragraph 3(c) on page 2, as follows:
"Spawning reaches - sections within the spawning rivers as
follows:
1. James River: from a line connecting Dancing Point
and New Sunken Meadow Creek upstream to a line
connecting City Point and Packs Point;
2. Pamunkey River: from the Route 33 bridge at West
Point upstream to a line connecting Liberty Hall and
the opposite shore;
3. Mattaponi River: from the Route 33 bridge at West
Point upstream to the Route 360 bridge at Aylett;
4. Rappahannock River: from the Route 360 bridge to
Tappahannock upstream to the Route 3 bridge at
Fredericksburg."
5.3.2 Alewife/Blueback Herring
Background. Participants agreed that the criteria
developed in this section would apply to the alewife, blueback
herring and other alosids. Alewife populations have declined
more than the blueback herring and are in greater need of
restoration. The group agreed this section should be rewritten
based on A.J. Lippson's compendium. Several documents
(Krauthamer and Richkus, 1987a, 1987b, 1987c, and 1987d) were
sources for the background narrative for the alewife.
Critical Life Stage. Both the egg and larval stages were
determined to be critical.
Critical Life Period. The beginning of the critical life
period was changed to "early March to the end of May."
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Food. The group agreed that food is critical, but asked
that the statement about larval feeding that appears in the
matrix be deleted.
Substrate. The substrate was determined to be not critical
for larvae, but critical for eggs and spawning since the
blueback herring has adhesive eggs.
Cover. SAV was determined to be not critical for larvae or
eggs.
Zone. This parameter was determined to be not critical.
Salinity. The group agreed that salinity is critical and
that the 0- to 5-ppt range in the Strawman II document was
acceptable.
Flow. Flow was determined to be not critical under natural
conditions, but important under conditions created by sheer,
power plant intake, pressure drop and dam turbines.
Temperature. The group agreed that temperature is a
critical parameter. The range of 16 to 24° C was determined to
be acceptable if it represents the lower and upper incipient
levels of larval response to temperature.
pH. The group agreed that pH is a critical parameter.
Members said the range of 6.5 to 8.5 appeared to be acceptable.
Dissolved Oxygen. Participants agreed that greater than
5.0 mg/1 was an acceptable criterion for dissolved oxygen for
the alewife but noted that this criterion would vary for
different species.
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Ionic Constituent. Not enough was known about this
parameter to determine whether it is critical.
Turbidity. Turbidity was determined to be critical.
Participants accepted the turbidity level of less than 50 NTU,
but noted that two-thirds of the population might show
decreased hatching success at this level.
Bacteria. The group noted that, although this variable was
related to water quality and anoxia, by itself it is not
critical for the alewife and herring.
Secchi Depth. This parameter was determined to be not
critical.
Suspended Solids. The group agreed that suspended solids
are critical to eggs. They changed the level to "50 mg/1."
Light Intensity. This parameter was determined to be not
critical.
Nitrogen and Phosphorus. Participants did not set any
levels for nitrogen and phosphorus, but requested that the
following points be noted: "Ammonia, nitrites and any form of
reduced nitrogen are known to be toxic. Nitrogen and
phosphorus can have direct toxic effects on finfish, but the
most critical impact is their collective affect on food
production and anoxia in stratified waters. These factors must
be taken into account when setting levels for these parameters
for finfish."
PAHs, Metals, Herbicides, Insecticides and Chlorinated
Hydrocarbons. These factors were considered to be
"provisionally critical." The Atlantic States Marine Fisheries
Commission Plan was referenced as a source for data on metals,
herbicides, insecticides and chlorinated hydrocarbons.
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Hardness and Alkalinity. Dr. Klauda (Johns Hopkins
University) may have data on hardness and alkalinity for the
blue herring, alewife and American shad. These data should be
used in the absence of species-specific data.
Geographic Distribution. Concerning geographic
distribution, the group recommended that Maryland's alosid
management plan, which describes all known spawning areas, be
used for distribution in Maryland. For all rivers with striped
bass, the distribution for alosids should extend from the lower
end of the spawning ground of the striped bass upstream to the
headwaters of all tributaries, except where fish would run into
a barrier, e.g., the West River and South River, in rivers in
which striped bass do not occur, the distribution of alewives
should be considered to go from the mouth of the river up to
any upstream blockage. These rivers are listed on the River
Herring Management Plan. The group recommended that
distribution of alosids in Virginia be based on the spawning
study by Dr. Loesch at the Virginia Institute of Marine
Science. The group also recommended that the distribution of
the alewife and herring as specified in the Pennsylvania
regulations be included.
5.3.3 Atlantic Menhaden and Spot
Critical Life Stage. Following the workshop, Dr. John
Merriner, National Marine Fisheries Service, Beaufort, North
Carolina, was contacted regarding the critical life stage of
the Atlantic menhaden. He said that the critical life stages
were eggs and larvae on the Continental Shelf and
post-metamorphic larvae and juveniles in the Chesapeake Bay.
Critical Life Period. The group accepted the critical life
period as being from April to October.
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Food. Food was determined to be critical, but the major
food items were not discussed (see also Chlorophyll on the next
page).
Cover. The group did not understand what was meant by
"shallow waters." Participants felt that cover is not critical
and queried whether the "shallow water" listing referred to the
larval or juvenile stage.
Zone. Zone was determined not to be a critical parameter.
The designation of zone was changed to "pelagic or open waters."
Salinity and Flow. These parameters were determined not to
be critical. "Estuarine" should be deleted.
Temperature. This parameter was determined not to be
critical. The limits were changed to "10 to 30° C."
pH. The group agreed this parameter is critical. They
accepted the 6.5 to 8.5 range given in the Strawman II document
and noted that the rate of change could affect survival. The
effect of acid rain on pH levels in the Bay should be
considered.
Dissolved Oxygen. Participants agreed this parameter is
critical. They accepted the greater than 5 mg/1 level given in
the Strawman II document as a minimal incipient level.
Ionic Constituent and Turbidity. These parameters are not
critical but can be lethal at extremes.
Bacteria. This criterion should be changed to pathogens.
An infectious pancreatic virus and fungal parasites were
mentioned as being pathogens of concern for the menhaden.
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Secchi Depth, Suspended Solids and Light Intensity. These
parameters are not critical because menhaden are found
naturally in turbid areas.
Chlorophyll. Chlorophyll is critical as food, but no level
was set. Phytoplankton cell size is critical, since menhaden
are unable to filter sizes less than 12 to 20 micrometers.
Nitrogen and Phosphorus. Participants did not set any
levels for nitrogen and phosphorus, but requested that the
following points be noted: "Ammonia, nitrites and any form of
reduced nitrogen are known to be toxic. Nitrogen and
phosphorus can have direct toxic effects on finfish, but the
most critical impact is their collective effect on food
production and anoxia in stratified waters. These factors must
be taken into account when setting levels for these parameters
for finfish."
Carbon. Participants agreed that particulate carbon (as
opposed to dissolved carbon) was tentatively critical as an
indicator of primary productivity (and their algal-based food
supply), and that it must be at a given level to sustain
populations. They changed the title of the parameter to
"Particulate Organic Carbon." This change should apply to all
finfishes.
PAHs, Metals, Herbicides, Insecticides and Chlorinated
Hydrocarbons. The Atlantic States Marine Fisheries Commission
Plan was referenced as a source for data on metals, herbicides,
insecticides and chlorinated hydrocarbons. The group deleted
the levels given in the Strawman II document for metals,
herbicides and insecticides.
Hardness and Alkalinity. These parameters are not critical.
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Chlorine. Chlorine was added to the list of critical
parameters. The group agreed that any amount of chlorine could
be detrimental to the species.
Geographic Distribution. The distribution as indicated on
the map that was supplied was incorrect. The menhaden is
ubiquitous unless constrained by stream size or behavior.
5.3.4 Other Species
Once matrices had been completed for the striped bass,
alewife/blueback herring, and Atlantic menhaden and spot, the
group moved quickly through the other species. They felt the
killifish would have many unique criteria. The hog choaker
would be related to habitat requirements for the spot. Bay
anchovies would be closely related to the menhaden responses to
habitat.
5.4 Conclusions
Participants recommended assigning a two- or three-person
team to each species. These teams would thoroughly research
the literature and fill out the matrices, with references for
each number. Then another workshop should be held to peer
review the criteria, with the team present to defend the
numbers.
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6. WATERFOWL/BIRDS PLANNING SESSION AND TECHNICAL WORK GROUP
6.1 Approach
The waterfowl/birds planning session and work group were
chaired by Dr. Matthew Perry, U.S. Fish and Wildlife Service
Patuxent Wildlife Research Center. The group divided the bird
species into three groups: ducks, wading birds and raptors.
Participants filled out matrices for the canvasback and great
blue heron and agreed that many of the criteria and comments
for these two species also applied to other ducks and wading
birds respectively. Some data were supplied for the redhead,
the black duck and the wood duck. The raptors - eagle and
osprey - were discussed separately. In assigning criteria, the
group tried to find levels that would be protective of at least
75% of the population. During the work group session, Dr.
Holland (Martin Marietta Environmental Systems) and Dr.
Stevenson (Horn Point Environmental Laboratories) were
consulted for information on benthic organisms and submerged
aquatic vegetation, respectively.
6.2 General Changes and Recommendations
The work group eliminated some parameters for some
species. The work group also changed some of the critical life
periods, especially breeding times for canvasbacks and redheads
in Canada. There was some discussion of what constituted a
critical life stage; i.e., should it be the most critical stage
during the time the birds are in the Bay area, or the most
critical stage in their life regardless of whether it occurs
while they are in the Bay. The group had difficulty discussing
the ecological parameters for the food items, because most
participants were not experts in these species. They
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recommended that such experts be present at any future
workshops. Participants were uncertain what was meant by the
terms "cover" and "zone" for bird species.
6.3 Waterfowl
Geographic Distribution. The group discussed whether
geographic distribution should include areas where the species
used to reside historically. They agreed that distribution
should include areas of importance in the 1950s and should also
list areas that are important in the 1970s and 1980s. The
management goal should be to establish conditions throughout
the 1950s distribution area that would make those regions
amenable to the species again.
The group agreed that the loss of submerged aquatic
vegetation in the upper Bay river systems had greatly reduced
these areas as feeding portions of habitats for waterfowl,
including the canvasback, redhead and black duck. Wood ducks
are found mainly in upper tributaries bordered by trees,
swamps, and marshes, and have been less affected by the loss of
SAV in the upper part of the Bay. The upper Bay and the upper
Potomac River are the areas of greatest concern for the ducks
and need immediate attention and restoration. Hope was
expressed for a return to pre-Agnes levels of SAV. The redhead
has almost disappeared from the entire Bay. Populations of
most duck species tend to be lower now than the high
populations of the mid-1950's due to the changing habitats of
the Bay.
Critical Life Stage and Period. The group decided that the
critical life stage for all ducks would be the adult stage when
they are wintering in the Bay. They felt that limited food
sources during this period made this stage more critical than
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nesting. In accordance with this decision, all critical time
periods for ducks were changed to "October through April."
Food. Some duck species have changed their food habits.
The black duck and canvasback are feeding more on molluscan
invertebrates. The redhead and widgeon have not changed their
food habits but have almost disappeared from the Bay due to
lack of SAV. Tundra swans and geese now feed on waste cereal
grains in agricultural fields rather than on SAV.
6.3.1 Canvasback
The correct species name is "canvasback/" not canvasback
duck. The matrices should reflect this change.
Critical Life Stage. The critical life stage was changed
to "wintering."
Critical Life Period. The critical life period was changed
to "October through April."
Geographic Distribution. Traditionally, canvasbacks fed in
the upper Bay early in the season and moved down the Bay as the
water froze. Stewart's research in the 1950s (Stewart, 1962)
indicates that the canvasback habitat covered the entire upper
Bay at that time. Since the 1950s, the canvasbacks have
wintered in the Susquehanna flats (historically), all eastern
shore tributaries north of the Choptank, the middle Potomac
north of Port Tobacco to Nomini Bay and Mobjack Bay.
Historically, the upper Bay was the most important area for
canvasback populations. Currently, the middle Bay (5 to 15 ppt
salinity zone) holds the most canvasbacks because of adequate
food reserves. A top priority for canvasbacks should be to
restore them to the freshwater areas of the Bay.
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Food. The group agreed that, historically, Vallisneria
americana, Potamogeton pectinatus and Macoma balthica were all
very important food species for the canvasback. Potamogeton
perfoliatus, Ruppia maritima, Zostera marina and Rangia cuneata
were of secondary importance. A survey of 323 canvasbacks from
1970 to 1979 (Perry, in press) showed that the canvasbacks1
diet has changed due to changing availability of food species.
In this survey, the predominant food of the 323 canvasbacks was
as follows:
85% - Macoma balthica
5% - Rangia cuneata
3% - Mya arenaria
1% - Leptocheirus plumulosus
1% - Nereis sp.
2% - Ruppia maritima
1% - Potamogeton perfoliatus
Myriophyllum spicatum is not a food source for canvasbacks and
should be deleted from the matrix. The group added crustaceans
(including mud crabs, arthropods and isopods) as an important
food species. They also tentatively added Corbicula
manilensis, an Asian freshwater clam that is present in the
Chesapeake Bay. Ducks eat these clams in Taiwan. It is not
known if they are an important food source in the Chesapeake
Bay. This could be an area for research.
Substrate. The group changed the substrate for Ruppia
maritima to "prefers sand or silty mud."
Cover. The group was uncertain as to what "cover" meant
for bird species. Members agreed that cover was not an
applicable requirement for the canvasback.
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Zone. The group was uncertain what "zone" meant in terms
of bird species. Participants thought the figure of less than
3 meters seemed correct, but did not have the expertise to say
so with certainty.
Salinity. The group changed the salinity criterion for
Zostera marina to "5 to 35 ppt" and for Ruppia maritima to "5
to 60 ppt." (Higher salinity than sea strength may result due
to evaporation in wetlands not inundated by daily tides.)
Participants were unable to determine whether the other
criteria were valid.
Temperature. The group expressed doubt about the accuracy
of the temperature criterion given for Potamogeton pectinatus,
Ruppia marit i ma, and Zostera marina. Dr. Holland said the
optimum range for the Rangia cuneata was 10 to 15° C.
pH. Some food species experience germination problems
below pH = 5. Participants thought 6 to 9 might be an
acceptable range for pH for the food species, but were not
sure. They also felt that pH might have an effect on Macoma
balthica and Rangia cuneata and, therefore, the designation of
"not limiting" for these two species in the Strawman II
document might be incorrect.
Dissolved Oxygen. The group felt that "greater than 5
mg/1" might be an acceptable criterion for the six food species
numbered 13.1 to 13.6 but were not certain of this. Members
inserted a criterion of "5+_l mg/1" for Macoma balthica, and
changed the text for Rangia cuneata to read "Needs oxygen to
live."
Ionic Constituent and Bacteria. These criteria were
determined not .to be applicable.
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Turbidity. The group inserted a criterion of less than 20
mg/1 for Vallisneria americanaf Potamogeton pectinatus,
Potamogeton perfoliatus, Zostera marina, and Ruppia maritimaf
and changed the text for Rangia cuneata to read "does well at
high turbidity."
Secchi Depth. For Vallisneria americana, Potamogeton
pectinatus, Potamogeton perfoliatus and Ruppia maritima, the
group recommended that secchi depth should not be less than the
depth to the bottom. They thought that secchi depth might not
be a critical requirement for Macoma balthica and Rangia
cuneata, but they were not sure.
Suspended Solids. The group referenced the Turbidity
criterion.
Light Intensity. Dr. Stevenson thought the criteria given
in the Strawman document for light intensity were low. He
recommended that they be checked to make sure that the values
given were for full saturation rather than half saturation. He
thought that the numbers given probably came from the EPA
Technical Synthesis Report, 1983, by Wetzel and Van Tine. If
not, he suggested this reference be checked for comparison with
the values given.
Chlorophyll. The group inserted a criterion of "less than
15 ug/1" for Vallisneria americana, Potamogeton pectinatus,
Potamogeton perfoliatus, Zostera marina and Ruppia maritima.
Dr. Holland commented that Macoma balthica does well at high
levels of organics and that chlorophyll may be limiting for
Rangia cuneata under conditions of low dissolved oxygen.
Nitrogen and Phosphorus. The general comment was made that
these two requirements must be considered together. Levels of
one affect species tolerance for the other. For freshwater
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species (Vallisneria americana and Potamogeton pectinatus)/
total dissolved nitrogen should be less than 1.4 mg/1 in
conjunction with phosphate levels of less than 0.003 mg/1. For
mesohaline species (Potamogeton perfoliatus, Zostera marina and
Ruppia maritima), total dissolved nitrogen should be less than
0.14 mg/1 in conjunction with phosphate levels of less than
0.01 mg/1.
PAHs. Oil in the Sea (National Research Council/ 1985) and
an EPA report (U.S. EPA, 1980) were mentioned as sources of
data on this parameter.
N/P Ratios. These were not discussed by the group.
Carbon. The carbon requirement was not discussed by the
group except for the comment that Macoma balthica and Rangia
cuneata do well at high carbon levels.
Metals. Dr. Holland stated that there is no evidence for
biomagnification of any metals other than mercury up the food
chain (Dillon, 1984).
Hardness and Salinity. The group specified a range of 10
to 30 ppt for Macoma balthica and 1 to 15 ppt for Rangia
cuneata.
Herbicides, Insecticides and Chlorinated Hydrocarbons.
These parameters were not discussed, except to say that SAVs
are tolerant of insecticide levels.
6.3.2 Redhead
Critical Life Stage. The critical life stage was changed
to "wintering."
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Critical Life Period. The critical life period was changed
to "October through April."
Geographic Distribution. Redheads currently winter around
the Tangier, Smith and South Marsh islands off the eastern
shore. Historically, they also resided in the same areas as
the canvasbacks (i.e., throughout vegetated areas of the Bay).
The participants agreed that they would like to see the redhead
restored to these areas.
Food. The group agreed that Vallisneria americana,
Potamogeton pectinatus, Potamogeton perfoliatus and Ruppia
maritima were important food species, but were less important
now than they had been historically. Participants considered
Zostera marina to be the most important food species for the
redhead at present. They deleted Myriophyllum pectinatus,
Macoma balthica and Rangia cuneata from the list. The group
noted that redheads also accidentally eat small snails attached
to the SAV.
Other requirements of the redhead were not specifically
discussed. However, the same SAV food species were listed for
the redhead as for the canvasback. Thus, criteria for these
species presented above under "Canvasback" also apply to the
SAV food species for the redhead.
6.3.3. Black Duck
Critical Life Stage. The group changed the critical life
stage to "wintering." Most black ducks breed in northern New
England and eastern Canada (especially the Maritime
provinces). Black duck populations are comparatively
insignificant in Chesapeake Bay as a proportion of the total
breeding population; nevertheless, they do have certain
breeding areas which should be protected.
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Critical Life Period. The group changed the critical life
period to "October to April."
Geographic Distribution. Historically/ there were large
concentrations of black ducks in the Susquehanna flats and the
eastern Bay region. Many habitats have been destroyed due to
various forms of development and erosion along much of the
eastern shore. Participants agreed that they would like to see
the black duck restored to these areas at its 1950s population
levels.
Food. The group agreed that Vallisneria americana,
Potamogeton pectinatus and Potamogeton perfoliatus were
important food species, but deleted Myriophyllum spicatum,
Zostera marina, Macoma balthica and Rangia cuneata from the
list. Participants were not sure how important Ruppia maritime
was to the black duck. The group added marsh plants to the
food species list and emphasized that these were a very
important food source for the black duck. Dr. Perry also
mentioned Melampus bidentatus (coffee snail) as a food source
(Grandy, 1972).
Cover. The group cited emergent marsh vegetation (Spartina
sp., Zizania aguatica, and Iva) and woody vines and shrubs as
important cover for the black duck. Hunting blinds and trees
also supply cover during breeding.
The group did not discuss other requirements for this
species. The Strawman II document provides life history notes
for the black duck.
6.3.4 Wood Duck
Critical Life Stage. The critical life stage was changed
to "wintering."
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Critical Life Period. The critical life period was changed
to "October through April."
Geographic Distribution. The population status of the wood
duck is reasonably good, but its habitats are in need of
protection. The wood duck lives in forested fresh parts of
most Bay tributaries on both the eastern and western shores
from the wetland/floodplains to the river fall line. The group
recommended that the wood duck be restored to 1950s
distribution and population levels.
Food. Following the workshop, Dr. Perry listed four
species as being major food species for the wood duck:
Peltandra virginica (arrow-arum), Sparganium eurycarpum (giant
burreed), Polygonum sp. (tearthumb) and Quercus sp. (oaks). He
also provided data on salinity, flow, temperature and pH for
these four species (see below).
Cover. Dr. Perry noted that cover is "needed for young."
Salinity. The salinity requirement for the four food
species is 0 ppt since they are freshwater species.
Flow. The flow requirement for all four species is "tidal
and nontidal."
Temperature. Temperature is not a limiting requirement for
any of the four food species.
pH. pH for all four food species should be "less than 7.0."
Nitrogen and Phosphorus. Dr. Stevenson felt that nitrogen
and phosphorus would not be limiting for the four SAV food
species.
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Other Requirements. No other requirements were discussed
for the wood duck or its food species.
6.4 Wading Birds
The group felt that populations of herons and egrets were
the same as they were in the early 1900s. Wading Birds (Sprunt
et al.f 1978) was mentioned as an information source. The
references in Strawman II were also cited.
Geographic Distribution. Great blue and green-backed heron
use the wooded tributaries for nesting areas, so geographically
these areas are important (see Geographic Distribution for the
great blue heron, below). The other herons and egrets use the
islands, mostly south of the Bay Bridge in the middle part of
the Bay. Smith island up through island complexes (Hooper's)
to the north are essential parts of the Bay for the island
nesters. From the Bay Bridge north, there are almost no herons
except the green-backed heron. The green-backed can be found
in small numbers on many islands and tributaries in the Bay.
Virginia has a large majority of its wading birds on the
Atlantic side in protected areas (especially south of
Chincoteague). Waders winter south of the Chesapeake Bay.
There has been some habitat loss for the herons due to
bulkheading and flooding of trees to create duck habitats, but
the group did not know how important this loss was.
6.4.1 Great Blue Heron
Background. The Strawman II background text says that the
minimum habitat for the great blue heron includes wetlands
within a "specified distance (e.g., 1 kilometer)" of a
heronry. One participant commented that a distance of 3 to 5
kilometers would be more suitable.
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Critical Life Stage. The group agreed that the critical
life stage is the nestling as indicated in the Strawman II
document.
Critical Life Period. The critical life period for the
great blue heron was changed to "May to July."
Geographic Distribution. The great blue heron has a
widespread distribution, with many at Poole's island, the
Aberdeen Proving Grounds, and the Chester and Wye rivers.
Upper tributaries and wooded swamps are important habitat areas
for the great blue. The largest group of great blues (about
750 pairs) is on Nanjemoy Creek in the Potomac Nature
Conservancy. Great blues are also found in Canoe Neck Creek,
the north shore of the Potomac (very important), the upper
portion of the Rappahannock, the upper Pocomoke on the Eastern
Shore, the central Bay (South Marsh Island, the Smith Island
complex, Tangier). Maryland has Critical Areas' guidelines of
1,000 feet riparian area for great blues and other waders. The
group felt that current populations of the great blue heron
should be maintained, but did not see a need to try to increase
the population.
Food. The group agreed that all three food species listed
- Menidia menidia, Fundulus heteroclitus and F. majalis - were
very important. They also noted that the great blue heron is
an extreme generalist and will eat many other kinds of food,
including perch, rats, frogs and snakes.
Toxicants. Dr. Erwin (Patuxent Wildlife Research Center)
felt that contaminants were not a problem, based on studies by
five or six researchers in the last 20 years. However, he
noted that there had been some local problems within five miles
of contaminant sources.
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Other Requirements. No other requirements were discussed
for the great blue heron.
6.4.2 Little Blue Heron and Green-Backed Heron
The green heron is now called the green-backed heron.
Food. Toads and frogs are an important food source for
these two species (Martin et al., 1951). No studies have been
conducted of the food habits of the little blue or green-backed
heron.
6.5 Raptors
6.5.1 Osprey
General. The osprey has recovered from DDT; however,
recent reports show that osprey reproduction is reduced in the
middle part of the Bay. Dr. Mitchell Byrd (College of William
and Mary), Dr. Paul Spitzer (Horn Point Environmental
Laboratories), and Mr. Jan Reese (St. Michaels, Maryland) were
mentioned as experts on the osprey.
Geographical Distribution. Over 90 percent of the Bay is
important for the osprey. There are some 1,500 pairs in the
Bay area. They can be found in all coastal areas but not in
deep water. They venture approximately 3 or 4 kilometers up
tributaries and possibly farther up the Potomac. They live at
least as far north as Miller's island. Dr. Byrd and Dr. Reese
can provide the limits of this habitat. Ospreys live on
navigational buoys and duck hunting blinds; however, these
structures have been decreasing in number. Ospreys breed
during the summer and winter further south.
-53-
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Food. The group agreed that Brevoortia tyrannus (menhaden)
is an important food source for the osprey. However,
commercial fishing of Brevoortia tyrannus has been increasing,
particularly that of smaller size fish eaten by ospreys. In
the last 5 to 7 years, researchers have seen nestlings fighting
for food. The group felt that Brevoortia tyrannus experts
should be involved with the aspect of the report dealing with
the ospreys.
Metals. Mercury in fish was mentioned as a possible
problem for the osprey. The group thought that Dr. Stan
Wiemeyer at the U.S. Fish and Wildlife Service Patuxent
Wildlife Research Center might have data on this.
Chlorinated Hydrocarbons. The group suggested that Dr.
Wiemeyer may also have data relevant to the effects of
chlorinated hydrocarbons on the osprey.
6.5.2 Bald Eagle
General. The eagle has recovered from DDT. Dr. J.D.
Fraser (Virginia Technical University, Department of Fisheries
and Wildlife Science, Blacksburg, Virginia), Dr. M. Byrd
(College of William and Mary, Williamsburg, Virginia), and Mr.
Keith Cline (Raptor Information Center, National Wildlife
Federation, Washington, D.C.) were mentioned as sources of
information on bald eagles. Two references for Bay eagles are
Bald Eagles in the Chesapeake: A Management Guide for
Landowners (Cline, 1975) and Andrew and Mosher (1982).
Geographic Distribution. The Bay area is a major resource
for bald eagles during the nonbreeding season. Birds from the
north (into Canada) and south (to Florida) and central Atlantic
states use the Bay and tributaries, important habitat features
-54-
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include fisheries, shoreline perches, and roost sites. The
Potomac River, Caledon State Park on the Potomac (about 50 to
75 birds in summer), Aberdeen Proving Grounds (more than 100
birds), the Blackwater National Wildlife Refuge (20 birds), and
the James River are important areas for the eagle. Bald eagles
need a 1,500- to 5,000-foot buffer zone between their nesting
area and development. Dr. Jim Fraser and colleagues are
studying distribution, habitat use, and disturbance.
Food. The group added dead ducks as a food source for the
bald eagle. This food source currently presents a problem
because hunters still use lead shot, which can poison eagles
that consume dead ducks. Lead shot will be illegal throughout
the United states by 1991.
Cover. The group was not sure how this requirement
pertained to the bald eagle, but they noted that the eagle
needs a wooded area including snags (which serve the dual
purpose of supplying nesting locations and observation points
for prey surveillance).
Zone, Temperature, Dissolved Oxygen, Ionic Constituent,
Turbidity, Bacteria. The group agreed that these parameters
are not applicable to the bald eagle.
Chlorinated Hydrocarbons. The group suggested that Dr.
Wiemeyer at Patuxent Wildlife Research Center may have data
relevant to the effects of chlorinated hydrocarbons on the bald
eagle.
-55-
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7. BLUE CRAB TECHNICAL WORK GROUP
7.1 Introduction
The blue crab work group was chaired by Dr. John
McConaugha, Old Dominion University. The group focussed on the
blue crab, but pointed out that there are many other
ecologically important crustacean species that participants did
not have time to address.
7.2 Critical Life Stage and Period
The participants concluded that all life stages of the crab
are important, in contrast with other species in which one life
stage is critically sensitive. Crab larval stages are
critical; however, they probably occur outside the Bay. The
pre- and postmolt stages are critical for Crustacea.
Therefore, protective habitats must be available to protect the
crab throughout its life. Other important factors include
availability of cover (SAV), metabolic mobilization of
toxicants, and increased risk of predators.
7.3 Background
Participants made several changes to the background text.
The first sentence of the third paragraph was changed to read
"All blue crab spawning occurs in Virginia waters." The last
two sentences of this paragraph were changed to "Most females
mate during the late summer season in July, August or
September, and hatching is delayed until the following summer.
A female may also produce two or more sponges of eggs later in
-56-
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the summer." The first sentence of the fifth paragraph now
reads "Juvenile crab migrations up the Chesapeake Bay and its
tributaries begin in August." The following paragraph was
inserted between the sixth and seventh paragraph of the text:
Molting is a major physiological event in the crustacean
life history. Brachyurans molt frequently during the early
juvenile stages (7-10 days). The periodicity decreases
with age and increased size. Because the premolt and
postmolt phases are periods of high metabolic activity, the
animal may be more susceptible to environmental stress
during these periods.
In addition, the group recommended that the background text
(particularly the reference to migration in the second
paragraph) be checked against key references. The publication
Synopsis of Biological Data on Blue Crabs (Callinectes sapidus)
(Millikan and Williams, 1984) provides an annotated
bibliography of major references for the blue crab.
7.4 Matrix
Food
Blue crabs are hardy and eat any scavengeable material.
However, the group concluded that food could be limiting under
some circumstances for blue crabs. Seasonal changes and
certain environmental conditions, such as low dissolved oxygen,
may affect benthic organisms by limiting the surface area of
their habitat. This reduced area may then affect crab
survival. Important food species for crustaceans include:
juvenile finfish, mysids, and %mall sand crabs. Low prey
density may result in cannibalism. More attention should be
paid to food web dynamics.
-57-
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Cover
The group agreed that SAV may be important cover for
juveniles and for molting crabs.
Zone
Crabs are found throughout the Bay/ but there is a
difference in distribution of males and females since the
females migrate toward the Atlantic to release their eggs,
Zone varies by season, life history stages and sex.
Salinity
Salinity is an important parameter for larval stages. The
group accepted the 2 to 21 ppt levels given in the Strawman
document for juveniles and adults.
Flow
The group felt that flow could have some long-term impact,
particularly on spawning stocks in the lower Bay and on larval
distribution and transport. Long-term alterations in salinity
patterns may affect distribution of spawning females. This
could alter larval distribution by changing the transport
system.
Temperature
Extreme cold temperatures such as freezing over the Bay may
increase juvenile mortality. The group discussed whether cold
-58-
-------
would have greater impact on animals that had not had adequate
food, but no consensus was reached.
£H
This parameter was not discussed by the work group.
Dissolved Oxygen
The DO level was changed to "greater than 2 mg/1," because
some participants suggested this was the level at which the
benthic community started to be affected by low DO. Low DO
could possibly restrict the available habitat of crabs, forcing
them into shallow waters where they would be more concentrated,
in which case available food would become limiting. Dissolved
oxygen is important when seiches occur, spreading
low-oxygenated water into shallow zones which endangers crabs
and other species. The group did not have good data on the
food web dynamics of this interaction, and thought it should be
investigated.
Herbicides, Insecticides and Chlorinated Hydrocarbons
The group considered all three pollutants together. There
is no good evidence that ecological levels of contaminants are
affecting mortality of blue crabs, contaminants may affect the
behavioral response of crabs to other ecological parameters.
Pollutants may also be important during the pre- and postmolt
stages. Energy reserves are mobilized during these stages, so
crabs would be exposed to the body burdens of the pollutants.
On the Eastern Shore where fields overlap into the marsh, there
may be some local effects due to high organophosphate
-59-
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concentrations in spring. This might be an appropriate area to
conduct a study of pesticide residues in the crab.
The group said that more data were needed on the
relationship between body burdens of pollutants in crabs and
fecundity/embryo survivability; e.g., do pollutants in the yolk
affect survivability and fecundity?
Other Factors
The group did not feel that most of the other factors
(light intensity, secchi depth, turbidity, ionic constituents,
suspended solids, chlorophyll, nitrogen, phosphorus, PAHs, N/P
ratios, carbon, metals, hardness, alkalinity) had any direct
impact on any critical stage in the blue crab; however, they
may have indirect effects through the food chain and behavioral
responses.
7.5 Geographic Distribution
The group suggested that any efforts to monitor the effects
of environmental conditions on the blue crab focus on the lower
Bay. This area includes the spawning grounds and provides the
initial habitat for larvae/juvenile recruits entering from the
shelf nursery grounds.
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8. REFERENCES
Andrew, J.M. and J.A. Mosher. 1982. Bald eagle nest site
selection and nesting habitat in Maryland. J. Wildl.
Manage. 46:383-390.
Atlantic Menhaden Management Board. 1986. Supplement to
the Atlantic Menhaden Fishery Management Plan. Fisheries
Management Report No. 8. Prepared for the Atlantic States
Marine Fisheries Commission, 1717 Mass. Ave., N.W.,
Washington, D.C.
Atlantic States Marine Fisheries Commission. 1985. Fishery
Management Plan for the Anadromous Alosid Stocks of the
Eastern United States: American Shad, Hickory Shad,
Alewife and Blueback Herring. Phase II in Interstate
Management Planning for Migratory Alosids of the Atlantic
Coast. Atlantic Fisheries Commission. 1717 Mass. Ave.,
NW, Washington, DC 20036. XVIII + 347 pp.
Bonn, E.W. et al. 1976. Guidelines for Striped Bass Culture.
ISBM 0-9-13235-20-2. American Fisheries Society,
Washington, D.C.
Buckler, D., P. Mehrle, L. Cleveland and J. Dwyer. In Press.
Influence of pH on the toxicity of aluminum and other
inorganic contaminants to east coast striped bass. Water,
Air and Soil Pollution.
Cline, K. 1975. Bald Eagles in the Chesapeake: A Manage-
ment Guide for Landowners. National Wildlife Federation,
Washington, D.C., 16 pp.
Davis, J., B. Laird et al. 1976. Effects of Tropical Storm
Agnes on the Chesapeake Bay Estuarine System. Johns
Hopkins University Press.
Dillon, T.M. 1984. Biological Consequences of Bioaccumulation
in Aquatic Animals: An Assessment of the Current
Literature. Technical Report D-84-2. Department of the
Army, U.S. Army Corps of Engineers, Washington, D.C.
Grandy, J.W. 1972. Winter Ecology of Maritime Black Ducks
(Anus rubripes) in Massachusetts, with Special Reference to
Nauset Marsh, Orleans and Eastham. Ph.D. dissertation,
University of Massachusetts, Amherst, 111 pp.
Holland, F. 1986. Long-term variation of macrobenthos in a
mesohaline region of Chesapeake Bay. Estuaries 8:93-113.
-61-
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Holland, F. et al. 1984. Long-term benthic monitoring for the
Maryland portion of Chesapeake Bay, July 1984-June 1986.
Prepared for Office of Environmental Programs, Department
of Health and Mental Hygiene, Maryland. Martin Marietta
Environmental Systems, Columbia, Maryland.
Krauthamer, J., and W.A. Richkus. 1987a. Maryland Department
of Natural Resources Management Plan for Alewife and
Blueback Herring. Draft. Prepared by Versar, Inc.
(formerly Martin Marietta Environmental Systems), Columbia,
Maryland 21045. March 1987.
. 1987b. Management Plan for Maryland's American
and Hickory Shad Stocks. Draft. Prepared for Tidewater
Administration by Versar, Inc. (formerly Martin Marietta
Environmental Systems). March 1987.
. 1987c. Characterization of the Biology of and
Fisheries for Maryland Stocks of Alewife and River
Herring. Source document. Draft. March 1987.
. 1987d. Characterizations of the Biology of and
Fisheries for Maryland Stocks of American and Hickory
Shad. Draft. Prepared for Tidewater Administration by
Versar, Inc. (formerly Martin Marietta Environmental
Systems). February 1987.
Martin, A.C., H.S. Zin, and A.L. Nelson. 1951. American Wild-
life and Plants. Dover, 500 pp.
Maryland Department of Natural Resources. 1986. Draft Final
Report on Polynuclear Aromatic Hydrocarbons and the
Chesapeake Bay. Maryland Department of Natural Resources,
Annapolis, MD.
Mehrle, P.M., L. Cleveland, and D.R. Buckler. In press.
Chronic Toxicity of an Environmental Contaminant Mixture to
Young (or Larval) Striped Bass. Water, Air and Soil
Pollution.
Mehrle, P.M. and L. Ludke. 1984. Early Life Stages of Striped
Bass, Progress Report 1980-1983. Columbia National
Fisheries Research Laboratory, U.S. Fish and Wildlife
Service, Columbia, Missouri.
Millikan, M. and A. Williams. 1984. Synopsis of
Biological Data on Blue Crabs (Callinectes sapidus). Food
and Agriculture Organization Fisheries Synopsis No. 138.
National Research Council, Ocean Studies Board. 1985. Oil in
the Sea, Inputs, Fates, and Effects. National Academy
Press, Washington, D.C.
-62-
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Perry, M. In press. Food habits and distribution of
wintering canvasbacks (Aythya vallisneria).
Setzler, E.M., W.R. Boynton, K.V. Wood, H.H. Zion, L. Lubbers/
N.K. Mountford, P. Pulles, and L. Tucker. 1980. Synopsis
of Biological Data on Striped Bass, Morone saxatilis
(Walbaum). NOAA Technical Report, Food and Agriculture
Organization Synopsis No. 121.
Sprunt, A., J.C. Ogden, and S. Winckler. 1978. Wading Birds.
Research Report No. 7 for the National Audubon Society, New
York, 381 pp.
Stevenson, C., and N. Confer. 1978. Summary of Available
Information on Chesapeake Bay Submerged Vegetation. U.S.
Dept. of the Interior, Fish and Wildlife Service,
F&W/OBS78/66.
Stewart, R.A. 1962. Waterfowl populations in the Upper
Chesapeake Region. Special scientific report. Wildlife:
65. U.S. Fish and Wildlife Service, Washington, D.C., 208
pp.
U.S. EPA. 1980. Ambient Water Quality for Polynuclear Aromatic
Hydrocarbons. EPA Report No. 440/5-80-069. NTIS PB
#81-117-806. Natl. Technical Information Service,
Springfield, Virginia.
Westin, D. and B. Rogers. 1978. Synopsis of Biological Data
on the Striped Bass, Morone saxatilis (Walbaum) 1792.
University of Rhode Island Marine Tech. Report 67.
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APPENDIX A
WORKSHOP AGENDA
-------
WORKSHOP ON HABITAT REQUIREMENTS FOR
CHESAPEAKE BAY LIVING RESOURCES
February 24, 1987
The Annapolis Ramada
Annapolis, Maryland
Sponsored by the
Chesapeake Bay Program's Living Resources Task Force
AGENDA
8:00 AM Registration and Chairmen Meeting
PLENARY SESSION
9:00 Welcome and Introduction
Living Resources Task Force Chair: Lee Zeni, Interstate Commission on
the Potomac River Basin
9:15 Overview of the Workshop Approach and Objectives
Workshop Chair: Maurice Lynch, Chesapeake Research Consortium
9:30 Concurrent Planning Sessions
• Benthos
Chair: Fred Holland, Martin Marietta Environmental Systems
• Plankton
Chair: Kevin Sellner, Benedict Estuarine Research Laboratory
• Submerged Aquatic Vegetation
Chair: Court Stevenson, Horn Point Environmental Laboratory
• Shellfish
Chair: Roger Newell, Horn Point Environmental Laboratory
• Finfish
Chair: George Krantz, Maryland Department of Natural Resources
• Waterfowl/Birds
Chair: Matthew Perry, USFWS - Patuxent Wildlife Research
Center
10:30 Coffee Break
10:45 Concurrent Technical Workgroups
• Marine Spawning Finfish/Anadromous Finfish
Chair: George Krantz, Maryland Department of Natural Resources
• Molluscan Shellfish
Chair: Roger Newell, Horn Point Environmental Laboratory
• Crabs
Chair: John McConaugha, Old Dominion University
• Waterfowl/Birds
Chair: Matthew Perry, USFWS - Patuxent Wildlife Research
Center
A-l
-------
12:00 PM Group Lunchecm
1:00 Reconvene Concurrent Technical Workgroups
• Marine Spawning Finfish/Anadromous Finfish
Chair: George Krantz, Maryland Department of Natural Resources
• Molluscan Shellfish
Chair: Roger Newell, Horn Point Environmental Laboratory
• Crabs
Chair: John McConaugha, Old Dominion University
• Waterfowl/Birds
Chair: Matthew Perry, USFWS - Patuxent Wildlife Research
Center
3:30 Coffee Break
CLOSING SESSION
Chair: Maurice Lynch
3:45 Chairmen present Summary Reports from the Concurrent Sessions and
Workgroups
5:15 Closing Remarks: Review of the Workshop
Proceedings Report and Continued Enhancement
of Habitat Objectives
Maurice Lynch, Chesapeake Research Consortium
5:30 Adjourn
A-2
-------
APPENDIX B
LIST OP PARTICIPANTS
-------
WORKSHOP ON HABITAT REQUIREMENTS FOR
CHESAPEAKE BAY LIVING RESOURCES
PLANNING SESSION PARTICIPANTS
February 24, 1987 - 9:30 a.m. to 10:30 a.m.
BENTHOS
Chair;
Dr. Fred Holland
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
ERG Rapporteur:
Jan Connery
Mr. Lowell Bahner
U.S. Environmental
Protection Agency
CSC/CBP
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
*Dr. Elizabeth Bauereis
Baltimore Gas & Electric Co,
Fort Smallwood Complex
P.O. Box 1475
Baltimore, MD 21201
301-787-5118
Dr. Daniel Dauer
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
**Mr. Charles Frisbie
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-974-3151
Dr. John Kraeuter
Baltimore Gas and Electric
Co.
Crane Aquaculture Facility
P.O. Box 1475
Baltimore, MD 21203
301-335-3011
Dr. Harriette Phelps
University of the District
of Columbia
Department of Biology
4200 Connecticut Avenue, N.W.
Washington, DC 20008
202-282-7364
Ms. Linda Schaffner
Geological and Benthic
Oceanography Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7366
Dr. Anna Shaughnessy
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
PLANKTON
Chair;
Dr. Kevin Sellner
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
ERG Rapporteur;
Betty C. Ford
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-l
-------
Dr. Raymond Alden
Applied Marine Research
Laboratory
Old Dominion University
1034 West 45th Street
Norfolk, VA 23508
804-440-4195
Dr. Ray Birdsong
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
Dr. Dave Brownlee
Academy of Natural Sciences
Benedict Estuarine Research
Laboratory
Benedict, MD 20612
301-274-3134
Dr. Larry Haas
Physical Oceanography
Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7248
Dr. Fred Jacobs
Coastal Environmental
Systems, Inc.
1829 Old North Point Road
Baltimore, MD 21222
301-288-0111
Ms. Gail Mackiernan
Maryland Sea Grant
H.J. Patterson Hall
University of Maryland
College Park, MD 20742
301-454-6420
Dr. Harold Marshall
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
*Mr. Larry Minock
Council of the Environment
903 9th St. Office Building
Richmond, VA 23219
804-786-4500
Dr. Kent Mountford
U.S. Environmental
Protection Agency
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
Dr. James Sanders
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
Mr. Robert Siegfried
Virginia Water Control Board
2111 N. Hamilton St.
Richmond, VA 23230
804-257-6683
FINFISH
Chair;
Dr. George Krantz
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-974-3558
ERG Rapporteur;
Ruth Thaler
*Mr. Ralph Abele
Pennsylvania Fish Commission
P.O. Box 1673
Harrisburg, PA 17105
717-657-4515
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-2
-------
Dr. Herb Austin
Biological and Fisheries
Science Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7322
Mr. Eric Barth
Virginia Marine Resources
Commission
2401 West Avenue
P.O. Box 756
Newport News, VA 23607
804-247-2200
Dr. Denise Breitburg
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
Mr. Jim Colvocoresses
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
404-642-7307
Dr. H. carlton Haywood
Interstate Commission on the
Potomac River Basin
Suite 300
6110 Executive Blvd.
Rockville, MD 20852
301-984-1908
Dr. Jim Hoff
Virginia Council on the
Environment
903 9th St. Office Building
Richmond, VA 23219
804-786-4500
Ms. Bess Gillelan
U.S. Environmental
Protection Agency
CSC/CBP
410 Severn Avenue, Suite 112
Annapolis, MD 21403
301-266-6873
Mr. Carl McMorran
Susquehanna River Basin
Commission
1721 Front Street
Harrisburg, PA 17102
717-238-0422
Dr. William Richkus
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
*Dr. James Thomas
NOAA
Estuarine Program Office
1825 Connecticut St., N.W.
Washington, DC 20235
215-673-5243
Dr. Robert Ulanowicz
Chesapeake Biological
Laboratory
University of Maryland-CEES
P.O. Box 38
Solomons, MD 20688
301-326-4281
*Mr. Lee Zeni
Interstate Commission on the
Potomac River Basin
Suite 300
6110 Executive Blvd.
Rockville, MD 20852
301-984-1908
WATERFOWL/BIRDS
Chair:
Dr. Matthew Perry
Patuxent Wildlife Research
Center
U.S. Fish & Wildlife Service
Laurel, MD 20810
301-498-0331
Living Resources Task Force Member
**Living Resources Task Force Staff
B-3
-------
Dr. Michael Erwin
Patuxent wildlife Research
Center
U.S. Fish & Wildlife Service
Laurel, MD 20810
301-776-4880
**Mr. Steve Funderburk
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21403
301-269-5448
*Mr. Glenn Kinser
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21401
301-269-5448
SHELLFISH
Chair:
Dr. Roger Newell
Horn Point Environmental
Laboratory
University of Maryland-CEES
P.O. Box 775
Cambridge, MD 21613
301-228-8200
ERG Rapporteur:
David Heffernan
Dr. Eugene Cronin
12 Mayo Avenue
Bay Ridge
Annapolis, MD 21403
301-267-6744
Dr. Bill Goldsborough
Chesapeake Bay Foundation
162 Prince George Street
Annapolis, MD 21401
501-268-8816
*Dr. Steve Jordan
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-269-3767
Dr. Romuald Lipcius
Biological and Fisheries
Science Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7330
Dr. John Mcconaugha
Department of Oceanography
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-4698
Dr. William Van Heukelen
Horn Point Environmental
Laboratory
University of Maryland-CEES
P.O. Box 775
Cambridge, MD 21613
301-228-8200
SUBMERGED AQUATIC
VEGETATION
Chair:
Dr. Court Stevenson
Horn Point Environmental
Laboratory
University of Maryland
P.O. Box 775
Cambridge, MD 21613
301-228-8200
ERG Rapporteur:
Carolyn Mulford
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-4
-------
**Mr. Richard Batiuk
U.S. Environmental
Protection Agency
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
**Mr. Bert Brun
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21403
301-269-5448
Dr. Robert Orth
Biological and Fisheries
Science Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7392
Ms. Ricky Price
U.S. Environmental
Protection Agency
CBP/Computer Sciences Corp.
410 Severn Avenue
Annapolis, MD 21403
*^Living Resources Task Force Member
Living Resources Task Force Staff
**
B-5
-------
WORKSHOP ON HABITAT REQUIREMENTS FOR
CHESAPEAKE BAY LIVING RESOURCES
TECHNICAL WORKGROUP PARTICIPANTS
February 24, 1987
10:45 a.m. - 12:00 p.m.
and
1:00 p.m. - 3:30 p.m.
CRABS
Chair:
Dr. John McConaugha
Department of Oceanography
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-4698
ERG Rapporteur;
Betty C. Ford
Dr. Raymond Alden
Applied Marine Research
Laboratory
Old Dominion University
1034 West 45th Street
Norfolk, VA 23508
804-440-4195
Mr. Lowell Bahner
U.S. Environmental
Protection Agency
CSC/CBP
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
**Mr. Charles Frisbie
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-974-3151
Dr. Bill Goldsborough
Chesapeake Bay Foundation
162 Prince George Street
Annapolis, MD 21401
501-268-8816
Dr. Romuald Lipcius
Biological and Fisheries
Science Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7330
Ms. Gail Mackiernan
Maryland Sea Grant
H.J. Patterson Hall
University of Maryland
College Park, MD 20742
301-454-6420
Dr. Harold Marshall
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
*Mr. Larry Minock
Council of the Environment
903 9th St. Office Building
Richmond, VA 23219
804-786-4500
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-6
-------
Dr. Robert Orth
Biological and Fisheries
Science Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7392
Ms. Linda Schaffner
Geological and Benthic
Oceanography Division
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
804-642-7366
Dr. Anna Shaughnessy
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
Dr. William Van Heukelen
Horn Point Environmental
Laboratory
University of Maryland-CEES
P.O. Box 775
Cambridge, MD 21613
301-228-8200
ANADROMOOS FINFISH AND
MARINE SPAWNING FINFISH
Chair:
Dr. George Krantz
Maryland Department of
Natural Resources
Oxford Laboratory
South Morris Extended
Oxford, MD 21654
301-226-5193
ERG Rapporteur;
Ruth Thaler
*Mr. Ralph Abele
Pennsylvania Fish Commission
P.O. Box 1673
Harrisburg, PA 17105
717-657-4515
Dr. Herb Austin
Biological and Fisheries
Science Division
Virginia institute of Marine
Science
Gloucester Point, VA 23062
804-642-7322
Mr. Eric Barth
Virginia Marine Resources
Commission
2401 West Avenue
P.O. Box 756
Newport News, VA 23607
804-247-2200
*Dr. Elizabeth Bauereis
Baltimore Gas & Electric Co.
Fort Smallwood Complex
P.O. Box 1475
Baltimore, MD 21201
301-787-5118
Dr. Denise Breitburg
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
Dr. Ray Birdsong
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
Dr. Dave Brownlee
Academy of Natural Sciences
Benedict Estuarine Research
Laboratory
Benedict, MD 20612
301-274-3134
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-7
-------
Mr. Jim Colvocoresses
Virginia Institute of Marine
Science
Gloucester Point, VA 23062
404-642-7307
Ms. Bess Gillelan
U.S. Environmental
Protection Agency
CSC/CBP
410 Severn Avenue, Suite 112
Annapolis, MD 21403
301-266-6873
Dr. Larry Haas
Physical Oceanography
Division
Virginia Institute of Marine
Science
Gloucester point, VA 23062
804-642-7248
Dr. H. Carlton Haywood
Interstate Commission on the
Potomac River Basin
Suite 300
6110 Executive Blvd.
Rockville, MD 20852
301-984-1908
Dr. Jim Hoff
Virginia Council on the
Environment
903 9th St. Office Building
Richmond, VA 23219
804-786-4500
Dr. Fred Jacobs
Coastal Environmental Systems
Inc.
2829 Old North Point Road
Baltimore, MD 21222
301-288-0111
Mr. Carl McMorran
Susquehanna River Basin
Commission
1721 Front Street
Harrisburg, PA 17102
717-238-0422
Dr. Kent Mountford
U.S. EPA
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
Dr. William Richkus
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
Dr. Kevin Sellner
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
*Mr. Robert Siegfried
VA Water Control Board
2111 N. Hamilton St.
Richmond, VA 23230
804-257-6683
Dr. Court Stevenson
Horn Point Environmental
Laboratory
University of Maryland
P.O. Box 775
Cambridge, MD 21613
301-228-8200
*Dr. James Thomas
NOAA
Estuarine Program Office
1825 Connecticut St., N.W.
Washington, DC 20235
215-673-5243
Dr. Robert Ulanowicz
Chesapeake Biological
Laboratory
University of Maryland-CEES
P.O. Box 38
Solomons, MD 20688
301-326-4281
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-8
-------
WATERFOWL/BIRDS
NOLLOSCAN SHELLFISH
Chair;
Dr. Matthew Perry
Patuxent Wildlife Research
Center
U.S. Fish & Wildlife Service
Laurel, MD 20810
301-498-0331
ERG Rapporteur;
Carolyn Mulford
**Mr. Bert Brun
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21403
301-269-5448
Dr. Michael Erwin
Patuxent Wildlife Research
Center
U.S. Fish & Wildlife Service
Laurel, MD 20810
301-776-4880
**Mr. Steve Funderburk
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21403
301-269-5448
*Mr. Glenn Kinser
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21401
301-269-5448
Ms. Ricky Price
U.S. Environmental
Protection Agency
CBP/Computer Sciences Corp.
410 Severn Avenue
Annapolis, MD 21403
Chair;
Dr. Roger Newell
Horn Point Environmental
Laboratory
University of Maryland-CEES
P.O. Box 775
Cambridge, MD 21613
301-228-8200
ERG Rapporteur;
David Heffernan
**Mr. Richard Batiuk
U.S. Environmental
Protection Agency
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
301-266-6873
Dr. Daniel Dauer
Department of Biological
Sciences
Old Dominion University
1054 West 45th Street
Norfolk, VA 23508
804-440-3595
Dr. Fred Holland
Versar ESM Operations
9200 Rumsey Road
Columbia, MD 21045
301-964-9200
*Dr. Steve Jordan
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-269-3767
Dr. John Kraeuter
Baltimore Gas & Electric Co.
Crane Aquaculture Facility
P.O. Box 1475
Baltimore, MD 21203
301-335-3011
**
Living Resources Task Force Member
Living Resources Task Force Staff
B-9
-------
Dr. Harriette Phelps
University of the District
of Columbia
Department of Biology
4200 Connecticut Avenue, N.W<
Washington, DC 20008
202-282-7364
Dr. James Sanders
Benedict Estuarine Research
Laboratory
Academy of Natural Sciences
Benedict, MD 20612
301-274-3134
Living Resources Task Force Member
**Living Resources Task Force Staff
B-10
-------
APPENDIX C
LIST OF LIVING RESOURCES TASK FORCE MEMBERS
-------
LIVING RESOURCE TASK FORCE MEMBERS
MEMBERS
LIVING RESOURCE HABITAT CRITERIA WORKSHOP
February 24, 1987
PHONE NO./
REGION
Mr. Ralph Abele
Pennsylvania Fish Commission
P.O. Box 1673
Harrisburg, PA 17105
Ms. Elizabeth Bauereis
Baltimore Gas & Electric Co.
Fort Smallwood Complex
P.O. Box 1475
Baltimore, MD 21201
Mr. Louis Bercheni
Bureau of Water
Quality Management
P.O.Box 2063
Harrisburg, PA 17120
Dr. Steve Jordan
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
Mr. Glenn Kinser
U.S. Fish & Wildlife Service
1825 Virginia Avenue
Annapolis, MD 21401
Mr. Larry Minock
Council of the Environment
903 9th St. Office Building
Richmond, VA 23219
Dr. Louis Sage
Academy of Natural Sciences
of Philadelphia
19th and The Parkway
Philadelphia, PA 19103
Mr. Robert Siegfried
VA Water Control Board
2111 N. Hamilton St.
Richmond, VA 23230
717-657-4515
Pennsylvania
301-787-5118
Maryland
717-787-2666
Pennsylvania
301-269-3767
Maryland
301-269-5448
Chesapeake Bay
Region
804-786-4500
Virginia
215-299-1110
Chesapeake Bay
Region
804-257-6683
Virginia
PLANNING/
TECHNICAL
WORKGROUPS
Finfish
Anadromous
Finfish
Benthos
Molluscan
Shellfish
Finfish
Anadromous
Shellfish
Molluscan
Shellfish
Waterfowl/
Birds
Plankton
Crabs
Plankton
Crabs
Plankton
Marine
Spawning
Finfish
-------
Mr. Charles Spooner
U.S. EPA
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
Dr. James Thomas
NOAA
Estuarine Program Office
1825 Connecticut St., N.W.
Washington, DC 20235
Mr. Lee Zeni
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-266-6873
Chesapeake Bay
Region
202-673-5243
Chesapeake Bay
Region
301-269-2926
Maryland
Chairman
Shellfish
Molluscan
Shellfish
Finfish
Marine
Spawning
Finfish
Finfish
Anadromous
Finfish
STAFF
Mr. Richard Batiuk
U.S. EPA
Chesapeake Bay Liaison Office
410 Severn Avenue
Annapolis, MD 21403
Mr. Bert Brun
U.S. Fish and Wildlife
Service
1825 Virginia Avenue
Annapolis, MD 21403
Mr. Charles Frisbee
Maryland Department of
Natural Resources
Tidewater Administration
Tawes State Office Building
Annapolis, MD 21401
301-266-6873
Chesapeake Bay
Region
301-269-5448
Chesapeake Bay
Region
301-974-3151
Maryland
Mr. Steve
U.S. Fish
Service
1825 Virginia
Annapolis, MD
Funderburk
and Wildlife
301-269-5448
Chesapeake Bay
Region
SAV
Molluscan
Shellfish
SAV
Waterfowl/
Birds
Benthos
Crabs
Avenue
21403
Waterfowl/
Birds
Waterfowl/
Birds
- 2 -
-------
APPENDIX D
ADDENDUM TO BENTHOS WORK GROUP REPORT
By Fred Holland
Versar, Inc., ESM Operations
(formerly Martin Marietta Environmental Systems)
-------
THE REPRESENTATIVE IMPORTANT SPECIES (RIS) CONCEPT
It is not feasible to define habitat requirements for all
biota inhabiting estuarine systems like the Chesapeake Bay.
This is because the Bay consists of a diverse array of species
that have a broad range of habitat requirements. Habitat
requirements not only vary from species to species, but also
vary from life stage to life stage within most species. In
addition, habitat requirements vary geographically and seasonally,
It is, however, generally possible to identify biota which,
because of their abundance, distribution, ecological roles
(e.g., food web linkage), or economic importance (e.g.,
commercially exploited species), are essential to, and/or
representative of balanced indigenous populations of shellfish,
fish, and wildlife. These target species or RIS can be used to
focus definition of habitat requirements, making the assumption
that if populations of these surrogate species are protected,
then other populations and the ecosystem are protected. Because
many RIS are near the top of estuarine food webs or are key
links in food webs, changes in their abundance or distribution
indicate system-wide alterations. However, for RIS to be
reliable surrogates of habitat requirements, they must be
selected carefully. For example, RIS selections should include
biota that are sensitive to specific water quality parameters
as well as biota that are representative of all major trophic
levels. RIS should be selected from at least each of the
following categories:
• Species sensitive to specific water quality parameters
(e.g., dissolved oxygen or specific pollutants)
• Species using the habitats of the Bay as a spawning
and/or nursery ground (e.g. , species spawning in
estuarine and freshwater habitats)
• Species of commercial and/or recreational value
• Species that are habitat formers and are essential to
maintaining important ecosystem functions (e.g.,
submerged aquatic vegetation)
• Species that are important linkages in the food web
• Species recognized as threatened or endangered
• Nuisance species likely to be enhanced by changes in
water quality or other habitat requirements.
D-l
-------
RIS should also be selected to include:
• Primary producers and zooplankton
« Benthos
• Forage fish
• Predatory fish
« Other vertebates.
Definitions and protection of habitat requirements of only
"celebrity" species may not adequately protect essential
ecosystem functions.
D-2
-------
BENTHOS
Importance of Benthos
The Chesapeake Bay is home to an active community of
organisms which live in association with bottom sediments.
This assemblage, collectively known as the benthos, includes
familiar organisms such as oysters, clams, and crabs, as well
as less familiar forms, including segmented and unsegmented
worms, small crustaceans, snails, and anemones. A large por-
tion (~75%) of the living and dead organic material in the
Chesapeake Bay water, including the plankton and decaying plant
material washed in from the watershed, settles to the sediment
surface and decays. This decaying material is the major food
source for benthic organisms. As benthic organisms burrow
through the sediments to obtain this food, they alter sediment
characteristics. In addition, as a result of burrowing and
feeding activities, a portion of the nutrients and other chem-
icals buried in the sediments are returned to the overlying
water. Recycled nutrients frequently contribute to excess
phytoplankton production and eutrophication, and recycled
chemicals can contribute to local toxic problems and degraded
water quality. The Chesapeake Bay is a nursery ground for many
commercially and recreationally important fish. While on their
nursery grounds, many of these fish feed almost exclusively on
the benthos. In conclusion, benthic organisms are a Representative
Important Trophic Group forming important links between primary
producers and higher trophic levels and are an integral part of
the Bay food web.
Salinity/Dissolved Oxygen
Salinity is the major natural environmental factor con-
trolling regional distributional patterns for the Bay benthos.
Differences in sediment characteristics and in the levels of
bottom dissolved oxygen concentration that occur from shallow
to deep habitats control local benthic distributions as well as
differences in benthic communities that occur from the upper
Bay to the lower Bay. Most of the lower Bay (i.e., downstream
of the Rappahannock River), and high salinity regions of lower
Bay tributaries, are characterized by a diverse mix of species,
including deep-burrowing, longer lived species. Most of the
upper Bay is, however, characterized by shallow burrowing,
highly-productive, short-lived species. The benthic species
assemblage occurring in the upper Bay is similar to that char-
acteristic of eutrophic or stressed environments.
D-3
-------
Types of Benthic Communities
Several major assemblages of benthic populations occur
along the Bay's salinity and sediment gradients. These are:
(1) a tidal freshwater assemblage (sand and mud), (2) a trace
salinity assemblage (sand and mud), (3) a low salinity estuarine
assemblage (sand and mud), (4) a high salinity estuarine sand
assemblage, (5) a high salinity estuarine mud assemblage, (6) a
marine sand and muddy-sand assemblage, and (7) a marine mud
assemblage. The tidal freshwater assemblage is limited to the
upstream portions of Bay tributaries. Aquatic earthworms,
called oligochaetes, and larval insects are numerically dominant
in this habitat. The trace salinity assemblage occurs in the
transition zone between tidal freshwater and estuarine habitats.
It is of greatest extent in the upper portions of the mainstem
Bay and the Potomac and James rivers/and is of limited extent
in smaller tributaries. A mix of freshwater organisms which
tolerate exposure to low salinity, and estuarine species which
tolerate exposure to freshwater are abundant in the trace
salinity habitat. The low salinity estuarine assemblage is
dominated by estuarine species. A few marine species that
tolerate exposure to low salinity also occur in lower salinity
regions of the Bay. The high salinity estuarine sand and mud
assemblages are distinct assemblages, each dominated by marine
species that tolerate exposure to low salinity. The marine
sand and muddy-sand assemblages occurs over much of the lower
mainstem Bay and consists mainly of deep-burrrowing polychaete
worms. Epifaunal organisms are frequently attached to the tubes
of some of these deep burrowing biota. Most of the species
inhabiting high salinity assemblages do not tolerate exposure
to low salinities. The marine mud assemblage mainly occurs in
deep channels of the lower Bay and near the mouths of lower Bay
tributaries. Polychaete worms also dominate this habitat.
Geographic Distribution
The spatial distribution of benthic biomass for the Maryland
Bay is summarized in Fig. 1. The height of the bars represents
the average annual amount*of benthic biomass per square meter
of bottom area. The deep central portion of the Bay and the
lower half of the Potomac River support the lowest benthic
biomass. Low benthic biomass also occurs in the deeper regions
near the mouths of smaller tributaries. In these habitats.
annual abundance and biomass of benthic organisms is depressed
because of adverse effects associated with oxygen-depleted
(i.e., anoxic) bottom waters that occur during warmer months.
The effects of anoxia on the benthos are most apparent just
downstream of the Bay Bridge where anoxia is generally most
severe and of greatest duration. Benthic organisms occurring
in habitats that experience anoxia are small, rapidly-growing
forms that can reproduce in any season.
2
*In gms ash-free dry wt/m ,
-------
CHESAPEAKE BAY
0 5 .0 NAUTiC&<_ MILES
Figure 1. Spatial distribution of average annual benthic
biomass in the Maryland portion of Chesapeake Bay.
Bars are average values*when multiple stations
occurred in a region. The shaded contour shows the
region affected during the summer by anoxic bottom
waters.
D-5
egms ash-free dry wt/m .
-------
(a)
0.5
f jpolycftaete diomass
Crustacean blomass
Mo Husk tJiomass
in
^>.
+j
x
>•
c.
D
at
C/1
VI
o
1-4
OJ
JAN MM APR NAT JUN JUL UJG SEP OCT 3cC
Figure 2. Seasonal filuctuations in benthic biomass for repre-
sentative habitats. (a) high salinity sand habitat,
(b) high salinity mud habitat, (c) low salinity
estuarine assemblage.
D-6
-------
Shallow habitats along the margins of the upper Bay and
the lower half of the Potomac River do not experience summer
anoxia and are characterized by much greater benthic biomass
than the adjacent deeper habitats that experience summer anoxia.
A variety of benthic organisms are abundant in shallow habitats
including small, rapid-growing polychaetes and larger, slower-
growing crustaceans and mollusks. These habitats are the
primary nursery grounds for juvenile fish. Most of the lower
Bay does not experience anoxic waters and benthic biomass in
these habitats is high throughout the year. These habitats
are also important feeding and nursery areas for fish and crabs.
The greatest biomass of benthos, represented by the tallest
bars in Fig. 1, occurs in trace salinity and low salinity
estuarine habitats. Much of the suspended sediment and organic
inputs to the Bay is deposited in this habitat. The Macoma
clam, Macoma balthica, and the brackish water clam, Rangia
cuneata, comprise most of the benthic biomass in the zone of
maximum turbidity. These clams are particularly well adapted
to feeding on micro-organisms associated with organically rich,
frequently resuspended sediments.
Seasonal Variation
The biomass of benthic organisms at any one place in the
Bay fluctuates as much or more over an annual cycle than
from place to place. Figure 2 summarizes month-to-month
variation for the benthos of typical Bay habitats. In all
habitats, peak benthic biomass occurs in the spring (Fig. 2).
Factors influencing within-year variation in benthic biomass
vary among habitats. Essentially no benthic organisms survive
anoxic conditions that occur in deep habitats during summer
(Fig. 2a). When anoxic conditions dissipate in early fall,
deep habitats are repopulated within weeks by small, rapidly
growing polychaetes. Benthic biomass is also low during summer
in shallow habitats along the margins of the Bay and its
tributaries. Summer low biomass values in shallow habitats
are, however, larger than peak biomass values in deep habitats
that experience anoxia (Fig. 2a and 2b). A variety of taxa
contributes to biomass peaVs in shallow habitats, including
polychaetes, crustaceans, and mollusks. Seasonal variation in
benthic biomass is reduced in the trace salinity habitat;
however, biomass levels in this habitat are always an order of
magnitude higher than those in other habitats.
Benthic Organisms as Water Quality Indicators
In the Patuxent River, the abundance of adult Macoma clams
peaked in 1978-1980 near the zone of maximum turbidity at the
D-7
-------
same time that suspended sediment and sewage loadings were at
the highest levels recorded for this system (Fig. 3). As dis-
cussed above, Macoma biomass is closely linked to the amount of
organic material that is produced within or input to the system.
Patuxent Macoma populations have declined since 1980 as suspended
sediment loadings have declined and as sewage treatment facili-
ties have been upgraded. Declining Macoma biomass indicates
that the amount of organic material accumulating in Patuxent
sediments is decreasing and overall water quality is improving.
These data suggest that pollution abatement and cleanup programs
for the Patuxent River are effectively improving water and
sediment quality by limiting inputs and production of organic
material. These trends are not, however, related to specific
changes in water quality parameters (i.e., reduced inputs of
pollutants), but are rather associated with overall improvements
in water quality (e.g., increased dissolved oxygen decreased
turbidity, reduced chlorophyll, etc.). The benthos are respond-
ing in a measurable and interpretable way to these improvements
and appear to be an early indicator of system-wide improvements.
Salinity
Natural effects of salinity fluctuations on long-term
benthic abundance trends are shown in Fig. 4 for the low salin-
ity estuarine assemblage from the middle reaches of the Potomac
River. This figure suggests that year-to-year fluctuation in
salinity during the reproductive periods is a major factor
influencing long-term trends for benthic organisms. Salinity
exerts the most influence over benthic distributions during
early life stages shortly after reproduction because these life
stages generally have narrower salinity tolerance ranges than
do adults. Long-term benthic responses to salinity and other
sources of natural variation (e.g., climate) can and must be
determined before benthic habitat requirements can be defined.
Synergism Among Parameters
Figure 5 summarizes the responses of an abundant Bay
benthic species, Macoma balthica, to temperature, salinity, and
the impact of man-induced pollutants. The response pattern
should be typical of that for most other Bay biota, including
fish and other benthos, and shows the complex interactions that
exist between natural water quality parameters and man-induced
water quality changes. If more than three natural and man-
induced factors had been included in the experiments shown in
Fig. 5, responses would have been more complex. This informa-
tion suggests that definition of specific habitat requirements
for estuarine biota is complex and that determination of values
for specific habitat criteria is impractical given present
-------
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0)
E
5 10 15
Temperature (°C)
Figure 5. The effect of temperature and salinity on median
survival time (h) of Macoma balthica at a chromium
concentration of 64 mg/l"1 (after Bryant, McLusky,
Roddie and Newberry, 1984)
D-10
-------
30000
D 20000
E
N
S
I
T
Y 10000-
•9
•8
A
6 L
I
N
5 I
T
0>JAN80 0IJAN81 0IJAN82 01JAN83 01JAN84
SAMPLING DATE
01JAN85 01JAN86
Figure 4. Long-term abundance pattern for the small crustacean.
Leptocheirus plumulosus, in the low salinity estuarine
region of the Potomac River. Note the relationship
between the magnitude of reproductive pulses and
salinity during the reproductive period.
D-ll
-------
knowledge. Rather, it is more practical to use this type of
information to identify habitat requirements and water quality
parameters to which the biota are particularly sensitive and
that can be influenced by resource management actions. It may
then be possible to define the important habitat characteristics
of the benthos (or other biota) that the ..esource manager can
strive to attain. It may then be possible to develop standards
that protect these sensitive habitat requirements.
Benthos as Water Quality Indicators
The composition of Bay benthic communities is determined
by ambient sediment and water quality. Therefore, the makeup
and abundance of organisms composing benthic communities are
likely to respond to changes in water and sediment quality
resulting from inputs of pollutants, pollution abatement pro-
grams, or other management actions taken to improve the Bay's
water quality. Because many benthic organisms live for 1-2
years, changes in their populations are an integration of
changes in environmental conditions occurring over their life
span and are frequently better indicators of water quality than
direct measurements. In addition, because benthic organisms
are relatively immobile, they complete their life cycle within
the Bay and often within specific regions of the Bay. Thus,
benthic responses to changes in water quality are likely to be
region specific and easily interpreted. Finally, as important
intermediate links in the Bay food web, benthic responses to
water quality changes are likely to be representative of the
responses of other living resources. The benthos are, there-
fore, good indicators of overall water quality and protection
of their habitat should ensure protection of most other Bay
biota.
Conclusions
• Benthic organisms are an important component of the Bay
ecosystem, serving as food for fish and crabs and
mediating exchange processes between bottom sediments
and the overlying water column. They should be consid-
ered a Representative Important Group for Chesapeake
Bay. Protection of the benthic habitat is essential to
maintenance of a balanced Bay ecosystem.
• Benthic organisms provide a sensitive indicators of
water and sediment quality that integrates over trophic
levels, over time, and over a number of important
environmental variables. Protection of the benthic
fauna should thus result in protection of many other
fauna.
D-12
-------
• The impact of low dissolved oxygen waters on bottom
habitats is difficult to measure directly but is clearly
evident in benthic communities.
• The long-term response of benthic organisms to reductions
in organic inputs and initial clean-up of the Patuxent
River has been documented and appears to be favorable.
• Benthic responses to pollution abatement can be accurately
tracked because natural sources of variation are known
and can be partitioned from responses associated with
pollution abatement and cleanup programs.
• Interaction between natural environmental conditions
and man-induced pollutants is complex and affects the
impact of pollutants in biota. These interactions must
be considered when establishing habitat requirements.
D-13
-------
APPENDIX E
ADDENDUM TO THE SHELLFISH PLANNING
SESSION AND TECHNICAL WORK GROUP REPORT
By John Kraeuter
Baltimore Gas and Electric Company
-------
APPENDIX E
ADDENDUM TO THE SHELLFISH PLANNING SESSION
AND TECHNICAL WORK GROUP REPORT
An important functional aspect of Chesapeake Bay is the
seasonal cycle. Freshwater input to the system in a pulsed
fashion is essential for maintaining oyster beds. Predator and
disease control on oyster-producing" areas is/ in part,
controlled by the wet and dry years experienced by the system.
It is important that this system behavior not be disrupted.
The fact that this varies in intensity from year to year is
vital to oyster production. Years of very strong spring flow
may push predators and diseases farther down the Bay. This
same process may kill small, diseased or otherwise weakened
oysters. If the process lasts long enough, it may eliminate an
entire year class. Mature oysters are more resistant to such
pulses. They maintain the integrity of the bed, so it is
better able to support the next few years' spatfall because of
lack of predation and disease. Because of these natural cycles
and lack of consistent spatfall, the farthest upstream bars
cannot support intense harvesting pressure.
Harvest pressure on oyster bars must be scaled to the
natural processes supplying oysters, nutrients and
environmental perturbations to the system. An appropriate
model for the freshwater input to the Bay would be the
historical salinity or freshwater inflow scaled by the river
system (with perhaps the exception of major events such as
Agnes). The point of using the records for each system is
that, although the entire drainage basin tends to act in much
the same way, a very intense pulse one year in the James River
may not be matched by that in the Susquehanna.
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The increase in nutrients in the spring brought about by
runoff is also important to the "scaling" of that year's
processes. While the Bay is now clearly too eutrophic,
management should reduce nutrient and silt input without
disrupting the pulses of freshwater and without disrupting the
natural cycles of nutrient pulses. An equal percentage
reduction in each time unit is preferable to a concentrated
effort in any one time unit. Space and time scale ecological
processes and are interrelated. Resource managers should be
very careful when manipulating such processes.
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APPENDIX F
GENERAL COMMENTS ON THE MATRIX APPROACH
TO DEFINING HABITAT REQUIREMENTS
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GENERAL COMMENTS ON THE MATRIX APPROACH
TO DEFINING HABITAT REQUIREMENTS
Several participants individually and collectively
commented on the matrix approach to resource management. These
comments are summarized here.
1. Users of the data - planners, etc. - should be present at
future workshops to provide guidance to scientists on the
kinds of data needed.
2. There were many errors in Strawman II that made scientists
uneasy about the quality of the final product.
3. All data in the matrices should be carefully referenced
and documented. References should be attached to the
report.
4. In order to fill out the matrices, a study team should
thoroughly research the literature and fill in the
matrices based on the best data currently available.
Then, a second workshop should be held to peer review the
numbers they have selected. The individuals who compiled
the data would defend the data at the workshop.
5. For matrices where the key species and food species are
different (e.g., birds that feed on plants), experts in
the food species should fill out the matrices rather than
experts in the key species.
6. The Strawman approach does not make allowances for
synergism among the parameters.
7. Key species should include all important species, not just
those that are endangered or politically important. Even
if a species is doing well now, we should know what
criteria are protective to guard against future changes
and threats to the species.
8. The terminology in Strawman should be clarified. For
example, what is meant by "substrate?"
9. It is difficult to put single numbers into the matrices
because these may change under different conditions.
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10. If standards are set based on target species, the needs of
species that were not examined may not be addressed. All
species should be examined.
11. Many of the matrix elements were irrelevant for some of
the key species.
12. For many species, several life stages or the entire life
cycle are critical.
13. The species designated as key need to be reevaluated.
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