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An Invitation to Monitor Florida’s Coastal Wetlands
COASTA L F LO R I DA
Adopt-A-WetlandTRAINING MANUAL
DRAFT EDITION, AUGUST 2015
This publication was supported by the National Sea Grant College Program of the U.S. Department of Commerce’s National Oceanic and Atmospheric Administration (NOAA), Grant No. NA 14OAR4170108. The views expressed are those of the authors and do not necessarily reflect the view of these organizations.
Additional copies are available by contacting Florida Sea Grant, University of Florida, PO Box 110409, Gainesville, FL, 32611-0409, (352) 392.2801, www.flseagrant.org.
SGEB 71 Draft, August 2015
DRAFTAUGUST2015
CoastalFloridaAdopt‐A‐WetlandTrainingManual
MaiaMcGuire1andLeRoyCreswell2
Editors
1FloridaSeaGrantAgent,UF/IFASExtension,FlaglerandStJohnsCounties
2RegionalAgent,FloridaSeaGrant
Florida Sea Grant
1762 McCarty Drive PO Box 110400
Gainesville, FL 32611-0400 (352) 392-5870
www.flseagrant.org
Take only pictures and leave only footprints.
Whenever on an adventure or working in the environment leave it like you found it.
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The Coastal Florida Adopt-A-Wetland program was adapted from the Coastal Georgia Adopt-A-Wetland
Program1 in 2015 to fit Florida’s dynamic coastal ecosystems.
This manual owes much of its success to the support, experience, and contributions of the following:
Mathew Monroe, University of Georgia Marine Extension Service, Georgia Sea Grant, Georgia Adopt-A-
Stream, Georgia Department of Natural Resources, Florida Department of Environmental Protection.
We are extremely grateful to all our volunteers for embracing the program and for all the good work they
are doing throughout the wetlands of coastal Florida. We would like to ask your assistance for the continued
improvement of this document. If you have suggestions for improvement of this manual, please send them
to Maia McGuire at [email protected].
1http://marex.uga.edu/wetland/
Acknowledgements
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Table of Contents
Acknowledgements ....................................................................................................................................................... ii
Chapter 1: Introduction ............................................................................................................ .................................. 1
How to Get Started ......................................................................................................................................... 2
How Will My Data Be Used? ......................................................................................................................... 4
Safety Issues ...................................................................................................................................................... 4
Coastal Wetland Habitats ................................................................................................................................ 7
Welcome to the Estuary .................................................................................................................................. 7
The Salt Marsh .................................................................................................................................................. 7
Salt Marsh Zonation ...................................................................................................................................... 10
Mangroves ....................................................................................................................................................... 12
The Beach ........................................................................................................................................................ 20
Chapter 2: Wetland Registration, Watershed Survey, and Map Assessment ..................................................... 24
Coastal Florida Adopt-A-Wetland Registration Form ............................................................................. 25
Coastal Florida Adopt-A-Wetland Watershed Survey & Map Assessment........................................... 27
How to Determine Your Latitude & Longitude ....................................................................................... 31
Chapter 3: Visual Monitoring ................................................................................................................................... 32
Visual Monitoring Protocol .......................................................................................................................... 33
Water Appearance .......................................................................................................................................... 33
Photo Documentation ................................................................................................................................... 33
Impaired Habitat Indicators ......................................................................................................................... 34
Wetland Condition / Appearance ............................................................................................................... 34
Soil Survey ....................................................................................................................................................... 34
Coastal Florida Adopt-A-Wetland Visual Survey Worksheet ................................................................. 35
Chapter 4: Biological Monitoring ............................................................................................................................. 37
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Biomonitoring................................................................................................................................................. 38
Diversity of Organisms in Estuarine Communities .................................................................................. 38
General Population Growth and Carrying Capacity in Natural Communities ..................................... 39
Density-Dependent vs Density-Independent Factors .............................................................................. 41
Comparison of r- and K-Selected Species .................................................................................................. 41
Diversity in an Estuary .................................................................................................................................. 42
Diversity of Estuarine Mud Flats ................................................................................................................. 44
Diversity of an Oyster Reef .......................................................................................................................... 45
How is Diversity Measured? ......................................................................................................................... 46
Monitoring Protocol ...................................................................................................................................... 47
Estuarine Bioassessment ........................................................................................................................ 47
D-Net Survey .................................................................................................................................... 47
Quadrat Survey ................................................................................................................................. 48
Hester-Dendy Survey ....................................................................................................................... 49
Beach Bioassessment .............................................................................................................................. 50
Hester-Dendy Survey ....................................................................................................................... 50
Seine Survey ...................................................................................................................................... 51
Mangrove Diameter and Height Measurement .................................................................................. 51
Coastal Florida Adopt-A-Wetland Biological Survey Worksheets
Coastal Florida Adopt-A-Wetland Biological Monitoring Form ..................................................... 53
Coastal Florida Adopt-A-Wetland Biological Community Sampling Form (Invertebrates) ........ 54
Coastal Florida Adopt-A-Wetland Biological Community Sampling Form (Vertebrates) ........... 55
Quadrat Survey Data .............................................................................................................................. 56
Cordgrass Height Data Sheet ................................................................................................................ 57
Shannon-Wiener Biological Diversity Index Worksheet ................................................................... 58
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Mangrove Diameter and Height Measurements ................................................................................. 59
Chapter 5: Physical/Chemical Monitoring ............................................................................................................. 60
Physical/Chemical Parameters ..................................................................................................................... 61
Temperature .................................................................................................................................................... 61
pH ..................................................................................................................................................................... 62
Soil & Sediment pH ....................................................................................................................................... 64
Dissolved Oxygen .......................................................................................................................................... 64
Salinity .............................................................................................................................................................. 66
Settleable Solids .............................................................................................................................................. 68
Turbidity .......................................................................................................................................................... 68
Physical/Chemical Monitoring Protocol .................................................................................................... 69
Physical Monitoring Protocol for Beach Site ............................................................................................. 73
Beach Slope Measurement: Emery Method for Beach and Dune Profiling ................................... 73
Longshore Current Measurement ......................................................................................................... 77
Florida Coastal Adopt-A-Wetland Physical/Chemical Survey Worksheets
Physical/Chemical Survey Water Monitoring Form ......................................................................... 78
Longshore Current Data Sheet ............................................................................................................ 79
Data Sheet for Beach Profiling ............................................................................................................ 80
Chapter 6: Problems in Your Adopted Wetland? ................................................................................................. 81
Wetland Watcher ............................................................................................................................................ 82
Dead or Dying Vegetation ............................................................................................................................ 82
Pollution .......................................................................................................................................................... 83
Marine Debris ................................................................................................................................................. 83
Derelict Traps ................................................................................................................................................. 84
Derelict Vessels .............................................................................................................................................. 85
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Microplastics ................................................................................................................................................... 85
Invasive Species .............................................................................................................................................. 86
Wildlife Violations ......................................................................................................................................... 88
Habitat Enhancement Projects .................................................................................................................... 88
“Who To Call” List ........................................................................................................................................ 89
Make Your Own Field Equipment .............................................................................................................. 90
Emory Rod Construction....................................................................................................................... 90
Make an Aquascope to Explore Tide Pools ........................................................................................ 91
How to Make a Viewscope .................................................................................................................... 92
Making a Quadrat .................................................................................................................................... 93
How to Make a Secchi Disk .................................................................................................................. 94
How to Make a Hester-Dendy Sampler ............................................................................................... 95
Make Your Own Plankton Net ............................................................................................................. 96
Sampling for Microplastics .................................................................................................................... 98
Bibliography ............................................................................................................................................................. 103
Appendices ............................................................................................................................................................... 104
Key to Macroinvertebrates Found in Coastal Florida ............................................................................ 105
Wetland Vegetation by Zones .................................................................................................................... 120
Florida Fish Identification Key .................................................................................................................. 130
Common Mollusks of Florida .................................................................................................................... 139
Other Common Marine Invertebrates of Florida ................................................................................... 142
Common Fishes of Florida ......................................................................................................................... 145
Introduced Non-native Aquatic Species in Florida ................................................................................. 148
Useful Websites ............................................................................................................................................ 149
Useful Books on Coastal Wetlands ........................................................................................................... 151
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HowtoGetStarted
HowWillMyDataBeUsed?
SafetyIssues
CoastalWetlandHabitats
Photo credits; Maia McGuire (top); US Fish & Wildlife Service (bottom)
ChapterOneIntroduction
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Welcome to the Coastal Florida Adopt-A-Wetland program! This is a hands-on education program that
promotes wetland conservation through volunteer monitoring. Wetlands are valuable coastal resources,
playing an important role in water quality, sediment retention, flood control, and wildlife habitat. This
program’s goals are namely to:
1. Increase public awareness of the state's nonpoint source pollution and water quality issues,
2. Provide citizens with the tools and training to evaluate and protect their local waterways,
3. Educate the public on the importance of wetlands, and
4. Collect quality baseline data and determine the health of our coastal wetlands.
HowtoGetStarted
This manual contains all the information you will need to begin monitoring your adopted wetland. This
manual is designed for groups that include school classes (5th grade through high school), civic organizations,
individuals, families, neighbors, friends, clubs, and companies. The first step is to attend a hands-on training
workshop where instruction will be provided on water quality monitoring and/or the biological-sampling
methods used to determine wetland habitat health. Volunteers who attend a training workshop will be
considered data collectors for one year. After an individual is certified in monitoring, a site must be chosen
for adoption and registered with their local Adopt-A-Wetland coordinator. A watershed survey and map
assessment must also be completed for this particular site on an annual basis. The watershed survey and
map assessment is a simple checklist of land uses and activities that influence your wetland site. If you have
questions or need help to complete the survey, call your AAW coordinator. It is also necessary to obtain a
map of the wetland you are choosing to adopt. This can easily be accomplished using Google Earth.2
2www.google.com/earth/
Introduction
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Many of the plants and animals that live in coastal wetlands are protected. When monitoring, be careful not
to damage vegetation. Remember, Take only pictures and leave only footprints. If collecting organisms to identify
back in the classroom, be sure to get the proper collection permits.3 Various levels of monitoring can be
conducted, ranging from basic visual surveys, which can be conducted multiple times per year, to monthly
water testing. Depending on your level of involvement, monitoring may consist of one or several of the
following:
Visual Monitoring: Participants conduct a simple visual survey multiple times per year,
consisting of observations of the plants, soil conditions, and water conditions.
Biological Monitoring: Biomonitoring determines the types and abundance of
vertebrates, macroinvertebrates, and plants that live in wetland areas. The diversity of species present helps
us to assess water quality and habitat health. Healthy ecosystems usually contain great diversity, and stressed
habitats support less species with a greater number of individuals (low diversity). This is also conducted
quarterly.
Physical/Chemical Monitoring: This involves the collection of information about specific water
quality parameters (e.g. temperature, pH, dissolved oxygen, salinity, turbidity, and settleable solids). This is
conducted on a monthly basis.
All levels of monitoring provide very important information concerning the health of coastal wetlands and
their protection. It is your decision as to which activities you will perform at your adopted site. This should
be decided based upon your group’s abilities and resources.
3http://fmsea.org/events/ascw/
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HowWillMyDataBeUsed?
Information acquired from the monitoring surveys will establish baseline data about the overall health of
your wetland. These data are valuable for verifying changes in your wetland over time and for identifying
potential pollution occurrences and/or die-off of any plants or animals. In addition to detecting problems,
data collected also will be used to establish baseline conditions for your area. Sometimes problems arise at
various wetland sites. Being aware of what to look for is important to the health of coastal wetlands. If
warning signs are spotted, volunteer groups should notify the appropriate agencies through their
“Emergency Contact List.”
SafetyIssues
Your safety is critical. Safety precautions need to be emphasized, especially on the coast. Sudden storms can
pop up and accidents can happen. The following information has been taken from the United States
Environmental Protection Agency’s website dealing with volunteer monitoring and assessing water quality.4
Follow these tips at your adopted site:
• Always monitor with at least one partner. Let someone else know where you are, when you intend
to return, and what to do if you do not come back at the appointed time.
• Develop a safety plan. Bring your cell phone or a radio. Locate the nearest medical center and write
down directions on how to get between the center and your site(s) so that you can direct emergency
personnel. Have each member of the sampling team complete a medical form that includes
emergency contacts, insurance information, and pertinent health information such as allergies,
diabetes, epilepsy, etc.
• Have a first aid kit handy. Know any important medical conditions of team members (e.g., heart
conditions or allergic reactions). It is best if at least one team member has first aid/CPR training.
• Listen to weather reports. Never go sampling if severe weather is forecast or occurs while at the site.
4http://water.epa.gov/type/rsl/monitoring/
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• Never wade in swift or rapidly rising water and watch for rip tides or currents.
• Watch for wildlife and insects such as ticks, hornets, and wasps. Know how to treat bites or stings.
• Watch for poison ivy, poison oak, sumac, and other vegetation that can cause rashes and irritation.5
• Do not monitor if the site appears to be polluted.
• Dress appropriately, with particular attention to your footwear.
• If at any time you feel uncomfortable about the condition of the wetland or your surroundings, stop
monitoring and leave the site at once. Your safety is more important than the data!
When using chemicals:
• Know your equipment, sampling instructions, and procedures before going out into the field.
Prepare labels and clean equipment before you get started.
• Keep all equipment and chemicals away from small children. Many of the chemicals used in
monitoring are poisonous. Tape the phone number of the local poison control center to your
sampling kit.
• Avoid contact between chemical reagents and skin, eye, nose, and mouth. Never use your fingers to
stopper a sample bottle (e.g., when you are shaking a solution). Wear safety goggles when
performing any chemical test or handling preservatives.
• Know chemical cleanup and disposal procedures. Wipe up all spills when they occur. Return all
unused chemicals to your program coordinator for disposal. Close containers tightly after use. Do
not switch caps.
• Know how to use and store chemicals. Do not expose chemicals or equipment to temperature
extremes or long-term direct sunshine.
• Be sure you have emergency telephone numbers and medical information with you at the field site
for everyone participating in fieldwork (including the leader) in case there is an emergency.
5https://edis.ifas.ufl.edu/ep220
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First Aid Kit
The minimum first aid kit should contain the following items:
• Telephone numbers of emergency personnel such as the police and an ambulance service.
• Several Band-Aids for minor cuts.
• Antibacterial or alcohol wipes.
• First aid cream or ointment.
• Several gauze pads 3 or 4 inches square for deep wounds with excessive bleeding.
• Acetaminophen for relieving pain and reducing fever.
• A needle for removing splinters.
• A first aid manual which outlines diagnosis and treatment procedures.
• A single-edged razor blade for minor surgery, cutting tape to size, and shaving areas before taping.
• A 2-inch roll of gauze bandage for large cuts.
• A triangular bandage for large wounds.
• A large compress bandage to hold dressings in place.
• A 3-inch wide elastic bandage for sprains and applying pressure to bleeding wounds.
• If a participant is sensitive to bee stings, include their doctor-prescribed antihistamine.
Think safe and be prepared:
Dressing for monitoring might not be fancy, but it works and will help keep you safe! Be prepared and use
common sense when working outdoors.
Wear shoes or boots, never go barefoot.
Do not forget your first aid, water and cell phone.
Be prepared for insects with repellent.
Remember your notebook and pencil.
When in the sun, a hat, sunglasses and long sleeves are a good idea.
Never go out alone/always go with a buddy and take your time, do not rush.
Oyster reef monitoring. Photo credit: Kay McGraw, NOAA
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WelcometotheEstuary
Estuaries are coastal wetland habitats where fresh water from land meets and mixes with salt water from the
sea. Ocean water contains approximately 35 parts per thousand (ppt, ‰) or 3.5% salt. Fresh water contains
0% salt. In an estuary, the salinity range is somewhere between these values. Florida has both saltmarsh
estuaries and mangrove swamp estuaries. Both are dynamic environments that are vital to the health of
Florida’s coasts and economy. Estuaries support a large and diverse population of marine life. They provide
habitat for juvenile fish. In Florida, nearly 90% of recreationally-targeted species and nearly 75% of
commercially-harvested marine species depend on estuaries during some portion of their life cycle.
TheSaltMarsh
CoastalWetlandHabitats
Photo credit: Holt/GTMNERR
8 DRAFTAUGUST2015
The salt marsh is characterized by expansive grasslands, mudflats, and meandering tidal creeks. Tidal creeks
flood and ebb, nourishing and flushing the salt marsh environment. Salt marshes are valuable because they
protect the mainland by absorbing the impact of storms. Salt marshes also help in filtering out harmful
pollutants that occur from a single location (point source pollution), or from a wide range of sources
(nonpoint source pollution) (Mitsch and Gosselink, 1986). People often ask how a wetland can absorb
toxins. Chemicals or pollutants may be absorbed into the mud, but may also be absorbed through the pores
of smooth cordgrass (Spartina alterniflora) and stored for a period of time within the plant’s body. (Long and
Mason, 1983).
Salt marshes and tidal creeks are important nursery grounds, providing a habitat for larval fish and shellfish
such as mullet, silverside, red drum, oysters and mussels. In addition, this productive and nutrient-laden
environment provides much needed organic matter for bacteria, which recycle the dead material into
valuable nutrients (Johnson et al., 1974) that act as fertilizers for phytoplankton (free-floating unicellular
plants). Many organisms depend on phytoplankton as a source of food, from the microscopic zooplankton
to the larger filter feeding animals including sea squirts, barnacles, clams, and mussels.
Oysters are also filter feeders and prominent members of the salt marsh community. They are described as
“keystone” species, which means they are critical in maintaining the health of this ecosystem. After
spawning, oysters begin their lives in the water column as free-floating larvae called “veligers”. When oysters
Photo credit: Oyster Restoration Working Group
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metamorphose, they settle and attach to a hard surface, becoming an immobile organism called a “spat.”
The spat or small oysters will often grow in clusters on other oysters in tidal creeks, on jetties, and on pilings
or docks. When oysters grow in clumps along tidal creeks they are called oyster bars or oyster reefs (O’Beirn
et al., 1994). If you closely examine the crevices of an oyster reef you, will find a variety of organisms, such
as mussels, crabs, fish, polychaete worms, and amphipods. Oyster reefs are called “Essential Fish Habitat”
which means that many commercially and ecologically important species depend on them in order to
successfully reproduce and survive. Oysters filter food particles such as plankton and detritus from the
water. Oysters are excellent at removing toxins, metals, nutrients, and harmful bacteria, thus improving
water quality. As it feeds, one healthy oyster can filter around 2.5 gallons (9.46 liters) of water through its
body in one hour. The more oysters we have in our creeks and estuaries, the cleaner our waters. Oyster reefs
also protect the marsh from being eroded by waves and boat wakes by absorbing the energy of these waves
before they can wash away the mud from around the roots of the Spartina grass.
Male fiddler crabs. Photo credit: Maia McGuire
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Other common organisms found in the salt marsh and associated mud flats are fiddler crabs, mud snails,
periwinkles, blue crabs, quahogs, shrimp, whelks, insects, killifish, mummichogs, wading birds, waterfowl,
and shore birds (Wiegert and Freeman, 1990).
The salt marsh can be divided into several zones. The low and high marsh zones will be our primary focus.
The low marsh is lowest in elevation and closest to the tidal creek and is the area that is flushed daily with
water during high tide. Smooth cordgrass (Spartina alterniflora) grows highest in areas that are closest to
creeks because of the frequent flushing of water when the tide floods. Plants that grow in the area furthest
from the tidal creek and at a higher elevation are classified as high marsh plants. Plants in this zone are
glasswort (Salicornia sp.), saltwort (Batis maritima), salt grass (Distichlis spicata), needle rush (Juncus roemerianus),
saltmeadow cordgrass (Spartina patens), sea oxeye daisy (Borrichia frutescens), marsh elder (Iva frutescens), and
eastern red cedar (Juniperus virginiana).
Because inundation of water occurs less often in the high marsh, salt tends to accumulate in the soil and it
becomes hypersaline, stunting the growth of Spartina alterniflora (Wiegert and Freeman, 1990). There are
other regions of the marsh where salt tends to accumulate. These regions are known as salt pans. Here the
concentration of salt in the soil is so high nothing will grow (Johnson et al., 1974).
Salt Marsh Zonation
Surprisingly, a large percentage of the mud and other soils in our coastal marshes come from areas inland. If
we traced the travels of a single sediment particle from the center of the state to the coast, the particle would
have a long adventurous trip. The particle begins inland as a part of a stream bank. Heavy rains will wash
out the bank of the creek causing the sediments to erode away into the water flowing downstream. Over
many years the sediment will venture down fresh water rivers as suspended material, and eventually enter
into brackish water (a mixture of fresh and salt water) found in estuaries.
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The silts and soils eventually fall out of the water column and accrete around saltmarsh grass and oyster
reefs. Suspended particles often fall out of the water column on the leeward or protected side of barrier
islands. This is why salt marsh formation generally occurs on the mainland side of Florida’s barrier islands.
The sediments build up mounds and piles that stick up above high tide. Smooth cordgrass (Spartina
alterniflora) eventually colonizes these areas. The accumulated mud with marsh grass begins its evolution into
a salt marsh. The processes of sediment build up (accretion) or sediment washing away (erosion) help shape
the salt marshes. Salt marsh mud is black; this is due to bacterial decomposition of organic materials which
causes a lack of oxygen in the soil and a chemical process called reduction. Reduction involves naturally-
occurring anaerobic bacteria, which are present in the mud. The bacteria have the capability to break down
one common seawater salt ion called sulfate (SO4) by using the oxygen and changing the sulfate into
hydrogen sulfide (H2S). This process is how the anaerobic bacteria utilize oxygen for respiration. All things
must respire in order to live. The rotten egg aroma prevalent in coastal marshes during low tide is the
hydrogen sulfide produced through this reaction.
The key to the importance of the estuary as an ecosystem lies in the plants that live and die there and the
bacteria associated with the mud. One important function of the marsh grass is to help hold the mud
together and to provide organic matter (detritus) to the coastal ecosystems. The bacteria in the mud and in
12 DRAFTAUGUST2015
the marsh utilize the detritus as a food source, recycling it into dissolved nutrients. Nutrients such as
nitrogen help with fertilizing the plants and algae, keeping the marsh system balanced and productive.
Mangroves
Mangroves are the predominant plant species of Florida’s central and southern coastal wetlands. Mangrove
swamps span some 2,700 kilometers of coastal Florida providing many ecosystem services that we as
humans depend on. These services, similar to those provided by salt marshes, including absorbing nutrients
and pollutants from runoff, protecting uplands from storm surge, and providing refuge for marine
organisms at all stages of life. Mangrove swamps are home to nearly 1,300 different species of flora and
fauna including endangered species and commercially valuable species.
A wide diversity of wildlife is typical in mangrove ecosystems. Florida mangroves are home to
approximately 220 fish species, 181 bird species (including the wood stork, white ibis, roseate spoonbill,
cormorant, brown pelican, egrets, and herons), 24 reptile and amphibian species (including alligators,
crocodiles, and turtles), and 18 mammal species (including bears, wildcats, pumas, and rats). Filter feeders
(especially barnacles, coon oysters and the eastern oyster) attach themselves to mangrove stems and prop
roots and filter organic material carried in by the tide. Crabs are another important mangrove species, as
they help maintain biodiversity. Crabs burrow in the sediments, prey on mangrove seedlings, facilitate litter
decomposition, and help convert detritus into energy forms that can be utilized by the ecosystem's birds and
fish.
With an area of about 600 square kilometers, Florida's Ten Thousand Islands is one of the world's largest
mangrove swamps. Much of its initial area was reduced by development, which led to mangrove
conservation laws.
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In Florida there are three species of mangrove: red mangrove (Rhizophora mangle), black mangrove (Avicennia
germinans), and white mangrove (Laguncularia racemosa). The buttonwood (Conocarpus erectus) is closely
associated with mangrove communities as well, but is not considered a true mangrove. Some mangrove
understories contain mangrove ferns, but few other plant species can survive the shady, high salinity
conditions.
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All mangroves have unique adaptations that allow them to live in estuaries. To survive in an estuary,
mangroves have adapted to the harsh environmental conditions such as anaerobic soil and salty water. Their
root adaptations are fundamental to their ability to survive. The roots give them the capability to anchor in
the sediment, conduct gas exchange, and help exclude salts. Mangroves are facultative halophytic species,
meaning they are plants that can grow in saline soil. There is little competition for mangroves because very
few plants can grow in these harsh, saline conditions.
Mangroves’ range in Florida is limited by climate. They thrive throughout the tropical and subtropical
regions of Florida. Northeast Florida and the Panhandle’s climate experiences occasional freezing
temperatures, which can stunt or kill mangroves.
Red Mangroves Rhizophora mangle
The name Rhizophora is derived from "rhizo" meaning "root" and "phora" meaning "carrier" or "bearer".
“Mangle” (pronounced main'-glee) is the Arawak Native American tribe’s name for the plant. Red mangroves
are easily identified by their tangled reddish aerial and prop roots. They have shiny dark green leaves which
are pale green on the underneath. Their flowers are yellowish-green. On average they do not exceed 30 feet
Aerial image of Ten Thousand Islands region of Florida. Photo credit: Everglades National Park
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tall, but individual trees can reach heights of over 100 feet tall. Red mangroves usually grow the most
seaward of the three species found in Florida and are the most susceptible to cold temperatures.
Red mangroves in south Florida. Photo credit: Maia McGuire
Red mangroves’ aerial and prop roots grow horizontally and vertically, allowing the trees to stand tall above
the anaerobic soil and laterally grow outward to deeper water. Small pores on the root surface, called
lenticels, allow oxygen to diffuse into the plant at low tide. Open passages in the roots, called aerenchyma,
move gases throughout the plant (Odum et al. 1982). Salt is excluded from the plant at the root surface;
excess salts in the plant are stored in the leaves and fruits.
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Red mangrove seeds go through continuous development and fully germinate on the tree. This process is
called vivipary, and the resulting structure is called a “propagule”. The propagule is a self-contained seedling
that grows between 12 and 18 inches long and looks like a long, curved cigar. When the propagule falls from
the tree, it may float for months before settling and rooting.
Black Mangrove Avicennia germinans
Named after the 10th-century Persian physician Avicenna, black mangroves were vital to early settlers of
Florida. Early Spanish settlers used the salty leaves for cooking. Later settlers made tea with the bark to treat
ulcers, hemorrhoids, and tumors. The branches were used to repel mosquitos; they would be placed in the
doorway and windows and allowed to smolder. Today, black mangrove flowers are utilized for production
of mangrove honey.
Red mangrove propagules on the tree. Photo credit: Maia McGuire
17 DRAFTAUGUST2015
Black mangroves are the least susceptible to cold temperatures.
Occupying slightly higher elevations than red mangroves, they grow
on average up to 60 feet, but can reach over 130 feet in height. Black
mangroves have aerial roots called pneumatophores that look like
pencils sticking out of the mud. Pneumatophores allow the plant’s
cable roots, which are buried in the anaerobic soil, to conduct gas
exchange.
The black mangrove’s bark is dark and scaly. The tree has clusters of
creamy white flowers, which produce seeds that develop into lima
bean-like propagules. Black mangrove propagules fully develop on
the tree before falling into the water and drifting to their final
destination. Black mangrove leaves have an elliptical, blunt shape
with a shiny dark green surface and a pale light green/gray underside
covered in dense hairs. The leaf blades have small salt glands that
excrete salt onto the leaf surface, often resulting in a visible
salty crust on the upper surface of the leaf.
White Mangroves Laguncularia racemosa
The white mangrove’s scientific name comes from a type of Roman jug called a "laguncula" which their fruit
resembles. White mangroves grow further inland than red or black mangroves, and are the smallest of the
three species in Florida. They reach heights of around 40 to 50 feet. When growing in anaerobic or stressed
soil, white mangroves may grow blunt-tipped pneumatophores similar to those of the black mangrove.
Black mangroves. Note the pneumatophores sticking out of the
sediment. Photo credit: Maia McGuire
Salt crystals on the surface of a black mangrove leaf. Photo credit: Maia McGuire
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The white mangrove has fleshy, flattened, oval shaped light green leaves with rounded ends. Two glands
located at the apex of the petiole excrete floral nectar from the plant. The flowers are greenish-white. The
propagules resemble small-flattened pears about an inch long. Like the red and black mangrove, the
propagules fall into the water after maturing. They drift until they are stranded, at which time small roots
will anchor them into the soil.
Buttonwood Conocarpus erectus
The buttonwood gets its name from the button-shaped fruit clusters it produces. Although not true
mangroves, because they lack mangroves’ reproductive and root characteristics, buttonwoods are closely
associated with them and are common in upper portions of mangrove swamps. Buttonwoods do not grow
as large as mangroves in Florida. They rarely reach heights of 15-20 feet and are often more shrub-like than
tree-like. The leaves are lance-shaped and have a silvery-gray sheen with two obliquely arranged nectar
glands. Unlike mangroves, the leaves are alternately arranged. Early settlers in South Florida cherished the
yellowish-brown heartwood of the Buttonwood—it was prized for fuel.
White mangrove leaves (left) and flowers (above). Photo credits: Maia McGuire (left),
NOAA (above)
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MangroveCommunities:Over-wash, Fringe, Riverine, Basin, Dwarf
Mangrove communities are characterized based on geomorphic and hydrological processes.
Over‐washmangrovecommunity
Over-wash communities are made up of small, low islands that experience frequent over-wash by tides. The
have high rates of organic matter.
Fringemangrovecommunity
Fringe communities make up swamps along canals, lagoons and rivers with limited tidal influences, where
shoreline elevation is slightly higher than mean high tide.
Riverinemangrovecommunity
Riverine communities make up the flood plains of tidal creeks. They produce high amounts of leaf litter.
Basinmangrovecommunity
Basin communities have low sheet flow and tidal influence.
Buttonwood tree showing the fruit that give the plant its name. Photo credit: Maia McGuire
20 DRAFTAUGUST2015
Hammockmangrovecommunity
Hammock communities have higher elevations in highly organic soil often surrounded by saltmarsh or other
types of marsh.
DwarfmangroveCommunity
Dwarf communities exist in South Florida and the Florida Keys. On average, heights only reach five feet. It
is believed the stunting of the mangroves is a result of low nutrient production in these regions and shallow
limestone substrate. All species of mangroves in Florida are dwarfed in these communities.
TheBeach
Florida’s thousands of barrier islands span over 1,260 miles of coastline. Florida has the longest coastline in
the U.S. except for Alaska (and is almost as long as the rest of the eastern seaboard of the U.S.). The beach
is another coastal “wetland” you may choose to monitor. Beaches are continually being shaped and changed
by ocean currents, waves, and wind. Coastal development, dredging, beach nourishment, and other human
impacts also play a role in shaping our beaches and dunes. The beach may be divided into three zones: the
shore (by the water’s edge), the wrack line (where decaying vegetation and other organic matter builds up),
and the sand dunes (mounds of sand held together by grasses and other plants.)
Sand dunes at Anastasia State Park. Photo credit: Maia McGuire
21 DRAFTAUGUST2015
Dune formation is a process dependent on the transport of dead vegetationfrom the estuary to the upper
beach area or wrack line. The dead vegetation deposited at the wrack line traps sand as it is moved by wind
and water. The mixture of the dead vegetation, sand, and moisture creates a soil mixture rich with organic
matter; suitable for plant growth. Seeds from salt-tolerant plants are deposited along the wrack line and
upper beach where they establish roots. The plants grow, trapping beach sand, and thus forming a primary
dune. In Florida, sea oats (Uniola paniculata) are the most important species of dune grasses because they are
mostly responsible for the creation of the primary dunes. Sea oats tolerate salty water, windy conditions, and
have the ability to thrive in harsh conditions. Sea oats build small sand dunes by trapping sand with the
expanding rhizomes, thus stabilizing loose sandy soil. These grasses are so valuable to Florida that they are
protected by the state, and any damage or removal can result in hefty fines.
As the primary dune evolves, it will either decrease in size from natural or man-made processes (erosion) or
increase in size (accretion) (Johnson et al., 1974). Over time, more plants will colonize and the dune will
eventually form a more diverse plant community, becoming a dune meadow, with plants such as pennywort
(Hydrocotyl bonariensis), yucca (Yucca aloifolia), camphorweed (Heterotheca subaxillaris), and dune primrose
Sea oat plants trapping sand. Photo credit: Maia McGuire
22 DRAFTAUGUST2015
(Oenothera humifusa). In addition, the dune system helps in providing a buffer for the mainland from the
ocean winds and storms.
Some of the animals commonly observed at the beach include ghost crabs, gulls, hermit crabs, jellyfish,
mole crabs, sea-whips, olive shells, whelk egg strings, sand dollars, horseshoe crabs, sponges, bottlenose
dolphins, and various fish including mullet, pompano and bluefish.In addition, five species of sea turtles are
found in Florida’s waters. Between May and August the threatened loggerhead sea turtle (Caretta caretta), the
endangered green sea turtle (Chelonia mydas), and at times the leatherback (Dermochelys coriacea) and Kemps
ridley (Lepidochelys kempi) sea turtles swim towards the coastal islands and beaches. These turtles leave the
water and dig a nest on the beach to lay 100-150 eggs. The fifth sea turtle species found in Florida waters is
the hawksbill (Eretmochelys imbricata). State and Federal laws protect all species of sea turtles.
Loggerhead sea turtle nesting. Photo credit: Ed Perry, Sebastian Inlet State Park
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The Florida Fish and Wildlife Conservation commission (FWC) asks the general public to report stranded
sea turtles by calling 1-888-404-FWCC (1-888-404-3922). If the sea turtle is tagged (with a plastic or metal
tag on a front or rear flipper), please include the tag color and number in the report if possible.
Never disturb a sea turtle that is crawling to or from the sea,
Observe nesting female turtles only from a distance,
Never attempt to ride a sea turtle,
Do not shine lights in a sea turtle’s eyes or take flash photography, and
Avoid or reduce beach lighting at night.
The coastal sand dunes, beaches, sandbars, and shoals comprise a vital natural resource system, known as
the sandsharing system. This system acts as a buffer to protect personal property and natural resources from
the damaging effects of floods, winds, tides, and erosion. Dunes and dune vegetation such as sea oats
(Uniola paniculata) are protected by law.
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RegistrationForm
WatershedSurvey&MapAssessment
HowtoDetermineYourLatitude&Longitude
ChapterTwoWetlandRegistration,WatershedSurvey&MapAssessment
Florida from space. Photo credit: NASA
25 DRAFTAUGUST2015
Coastal Florida Adopt-A-Wetland Registration Form
Complete the following form for each wetland you monitor and return to your local AAW coordinator. This form is to register a (circle one or more than one if necessary (e.g. beach and marsh) MANGROVE SALTMARSH BEACH
Latitude: Longitude:
Group Name:
Official Name of Site You Are Monitoring:
Level of Monitoring (Circle One/More): Chemical Biological Visual
Lead Coordinator/Contact: Today’s Date:
Complete Mailing Address:_______________________________________________
Phone Number(s):_____________________E-mail
Address:_____________________
For Official Use Only
Group has biological kit Group has chemical kit
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AAW Registration Form page 2
1. Describe the location of your monitoring site (e.g. 30 feet downstream of Arpieka Ave. and Inlet Dr. on Anastasia Island).
2. What is the name of your monitoring group? (e.g. Scout Troop 101, Friends of GTMNERR, Guana Lake Gators)?
3. What are the goals you hope to accomplish with the Adopt-A-Wetland program?
4. What equipment or supplies do you need to achieve your goals?
5. Where will you send the data you collect?
6. Name the data collectors in your group.
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COASTAL FLORIDA ADOPT-A-WETLAND Watershed Survey
To be conducted at least once a year and returned to your local AAW coordinator.
Adopt-A-Wetland Group Name:
Investigator(s):
__________________________________________________________________________
__________________________________________________________________________
Water Body Name:_________________________County(ies):
Picture/photo documentation? Yes No
Date: Time:__________________________
I. CREATE A MAP OF YOUR WETLAND
You can download a map from www.google.com/earth/ See instructions on page 31.
II. LAND USES/ACTIVITIES AND IMPERVIOUS COVER
a. Comments on general water body and watershed characteristics: (e.g. date and size of fish kills, increased rate of erosion evident, litter most evident after storms). Fish kills should be immediately reported to FWC 1-800-636-0511 or http://myfwc.com/fishkill or 888-404-FWCC (888-404-3922).
b. Summarize notable changes that have taken place since last year (if this is your second year conducting the Watershed Survey).
c. Identify land uses and activities near your monitoring site, which have the highest potential to impact water bodies:
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Check all boxes that apply, describe the location of the activity(ies) under the Notes on Location & Frequency of Activities and also mark the locations on your map. If you do not know some of the information below, write NA under Notes. Land Disturbance Notes on Location & Frequency of Activity
Erosion caused by land development or construction
Docks, piers, jetties
Large or extensive gullies
Unpaved roads near or crossing streams
Commercial forestry activities including harvesting and site-preparation
Extensive areas of creek bank failure or channel enlargement
Agricultural Activities
Croplands
Pastures with cattle access to water bodies
Confined animal (cattle or swine) feeding operations and concentration of animals
Animal waste stabilizations ponds
Poultry houses
Highways and Parking Areas
Shopping center & commercial areas
Interstate highways and interchanges
Major highways and arterial streets
Other extensive vehicle parking areas
Mining
Quarry
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Leisure Activities
Golf course
Marina(s)
Recreational Fishing
Boating/Jet skis
Swimming
Other
Transportation and Vehicle Services
Truck/car cleaning services
Automobile repair facilities
Auto dealers
Rail or container transfer yards
Shipping or fishing port
Marinas with boat fuel/repair/painting
Business & Industry, General
Exterior storage or material exchange
Activities with poor housekeeping practices indicated by stains leading to creek or storm drains or on-site disposal of waste materials
Heavy industries such as textiles & carpet, pulp & paper, metal & vehicle production
Dry cleaners or outside chemical storage
Special Issues
Fertilizer production plants
Feed preparation plants
Meat and poultry slaughtering or processing plants
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Construction Materials
Wood treatment plants
Concrete and asphalt batch plants
Waste Recycling, Movement & Disposal
Junk and auto salvage yards
Solid waste transfer stations
Landfills and dumps (old & active)
Recycling centers
Illicit Waste Discharges*
Sanitary sewer leaks or failure
Overflowing sanitary sewer manholes due to clogging or hydraulic overloading
Bypasses at treatment plants or relief valves in hydraulically overloaded sanitary sewer lines
Domestic or industrial discharges
Extensive areas with aged/malfunctioning septic tanks
Dry-weather flows from pipes (with detectable indication of pollution)
Signs of illegal dumping
* If found (most likely during watershed surveys), these activities should be immediately reported to the local
government or the DEP regional office.
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How to Determine Your Latitude and Longitude
You will need to know how to determine the latitude and longitude of your site so that others can find the
exact location. Google Earth is a great resource for determining your latitude and longitudeas well as for aerial
images of your adopted site. You can download Google Earth at no cost onto a desktop or laptop computer. In
Google Earth, you can enter your general location in the search box and then manually zoom in to your specific
location. The latitude and longitude coordinates for the location of the cursor will be at the bottom right of the
screen.
Many apps are also available for finding your latitude and longitude using an Apple or Android smart device.
Handheld GPS devices can also be used to determining latitude and longitude, and for mapping your wetland
site. Print a map of your site with the latitude and longitude of the area you will be monitoring. Keep a copy for
your group and send a copy to your Adopt-A-Wetland coordinator.
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MonitoringProtocols
VisualSurveyWorkSheet
ChapterThreeVisualMonitoring
Photo credit: Steve Davidson
33 DRAFTAUGUST2015
Visual monitoring represents the first level of activity in Coastal Florida’s Adopt-A-Wetland citizen science
monitoring program. The activities are basic, but are also very important. As you become the “Wetland
Watcher,” it is through your eyes that we can see problems that may occur in our valuable coastal wetlands.
Visual surveys are usually performed multiple times per year in a healthy wetland. However, when
monitoring an impaired or dead wetland site, we encourage you to perform the survey monthly. Remember
to always conduct the visual survey at the same stage of the tidal cycle each time if possible.
Monitoring Protocol
This protocol provides directions on performing a visual survey. Use the worksheet on pages 35-36 to
record important information about vegetation, soils and hydrology in your wetland.
Water Appearance
Fill a clean, clear container with water from your adopted wetland site. Hold the container up to the sun and
determine the color. Odor should also be easy to detect from the container, but sometimes you will not
notice an odor at all.
Photo Documentation
When monitoring a healthy wetland you should take a photograph of your site every visit. However, since
changes may occur rapidly, please include a photo each month when monitoring a marsh that appears
stressed. Please try to take the photo at the same spot and in the same direction each time. If possible also
take the photograph at the same tidal cycle (i.e. low or high tide).
VisualMonitoring
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Impaired Habitat Indicators
Sometimes human activities nearby will adversely affect a wetland. Check all the boxes in the impaired
habitat indicator section of your survey sheet that apply to your site.
Wetland Condition/Appearance
In this context, wetland condition refers to the health of specific plants that grow in the wetland. Marsh
grass, mangroves, and other plants naturally go through seasonal changes. The general pattern is that the
marsh grass turns brown in the winter but greens with new growth during the spring and fall. We expect
browning in the winter, however if the color stays brown year-round, we may have reason to become
concerned. Hard freezes may stress or kill mangroves; leaves will blacken and fall off the plant. Also pay
close attention to an abundance or absence of organisms (snails, crabs, or fish). If your adopted site shows
characteristics of continued stress please make your local Sea Grant Extension agent aware.
Soil Survey
The presence of large areas without plants (except for salt pans) may be a warning sign that something is not
right. These areas of the marsh will consist mostly of mud. If possible, and with permission, mark the
perimeter of the muddy area with PVC, sticks, or flags, or use a GPS and plot the perimeter of the marsh.
Each time you monitor your site note whether the muddy area devoid of plants is increasing or decreasing.
Dip your finger into the mud or scrape the surface of the sediment in your marsh and observe
characteristics of the sediment/mud. Check all the appropriate boxes on your survey sheet.
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Coastal Florida Adopt-A-Wetland Visual Survey
AAW Group Name County
Group ID Number Site ID Number
Investigators
Wetland Name Date Time
Picture/Photo Documentation? Yes No
Amount of Rain Inches in Last Hours/Days □Heavy Rain □Steady Rain □Intermittent Rain
Present Conditions: □Heavy Rain □Steady Rain □Intermittent Rain
□Partly Cloudy □Overcast □Clear/Sunny
Site Description: (e.g. Mangroves, Salt Marsh, Beach, Estuary)
Is Waterway Influenced by Tides? Yes No
If Yes, Tide was: □High □Outgoing □ Low □Incoming
Water Surface: □Calm □Ripples □Waves □Whitecaps
Impaired Habitat Indicators: □Foam □Bubbles □Oil □Scum
□Dead Organisms □Trash Present
□Dumping □Dredging
□Erosion □Vegetative Debris
□Excessive Algae Dock/Pier Present
□Artificial Water Control (groin, jetty, dyke, etc.)
Water Color: □Clear □Muddy □Milky Gray □Green
□Brown □Tan □Other
Odor: □Gas □Oil □Chlorine □Rotten Eggs
□Sewage □Chemical Other
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Wetland Condition/Appearance:
□Marsh Grass Green □Marsh Grass Brown □Other
□Marsh Surface Mostly Mud/ If so is mud area increasing or decreasing?
□Mangroves healthy □Mangroves dying □Other
Invertebrate Survey:
□Many White Snails (Periwinkles) □Few Periwinkles
□Many Black Snails (Mud snails) □Few Mud Snails
□Many Crabs □Few Crabs (What type of crabs?)
□Dead Fish Present □Dead Blue Crabs Present □Other Dead Organisms
□Ribbed Mussels Present □Dead Ribbed Mussels
Mud/Soil Survey:
□Black on Surface □Reddish Brown Color □Brown on Surface
□Black Surface Streaks □Mostly Brown Color □Green on Surface
□Other
Mud Moisture Content:
□Totally Dry □Wet □Damp
Mud Texture: □Clay/Mud (sticks to finger) □Sand (larger particles, not stick to
finger)
Additional Comments/Observations:
Submit form to your local Adopt-A-Wetland coordinator.
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Biomonitoring
DiversityofOrganismsinEstuaries
PopulationGrowthandCarryingCapacity
BiologicalMonitoringProtocolsandBioassessment
BiologicalSurveyWorksheets
Photo credit: GTMNERR
ChapterFourBiologicalMonitoring
38 DRAFTAUGUST2015
Biomonitoring provides information on changes in the plant and animal communities that may occur in our
wetlands. Any changes to these environments will be reflected in the quantity/quality of plants or the types
of animals present. There are various biological survey methods available to monitor your adopted site
depending on the type of site you may have.
Our monitoring protocol concentrates on macroinvertebrate (large visible animals without a backbone) and
vegetative monitoring. Some common types of macroinvertebrates include oysters, mussels, snails, crabs,
and worms. Some common plants, which may be found at your site, include mangroves, smooth cordgrass,
needle rush, sea oxeye daisy, and sea oats. Macroinvertebrates and wetland plants are good indicators of
wetland quality because:
• They are affected by the physical, chemical, and biological conditions of the wetland.
• They cannot escape pollution and show effects of short and long-term pollution events.
• They are an important part of the food web, representing a broad range of trophic levels.
• They are relatively easy to collect and identify with inexpensive materials.
Diversity of Organisms in Estuarine Communities
Biodiversity is a measure of the diversity or the number of different species occurring in a community. A
community is a naturally-occurring group of different species of organisms that live together and interact as
a unit. An estuarine community may contain animals such as the fiddler crab, marsh periwinkles, coffee bean
snails, clams, mussels, mud crabs, and stone crabs. Species diversity depends on species richness and species
evenness. Species richness is the number of species present, while evenness refers to the distribution of
individuals among the species (i.e. if all species are equally abundant then evenness is high, if a few species
are far more abundant than the rest then evenness is low).
Biomonitoring
39 DRAFTAUGUST2015
A high diversity would mean that there are many equally abundant species whereas a low diversity would
indicate few equally abundant species or many species with an unequal abundance. However, there are other
factors, which affect the numbers of species in coastal and marine habitats. General animal population
dynamics influenced by environmental factors and species strategies are inherent in nature. We have broken
down some of these population factors and the effects on the diversity of the coastal communities.
General Population Growth and Carrying Capacity in Natural Communities
The population growth of a species relates to an organism’s reproductive potential and environmental
factors within the habitat. Typically, population growth occurs in an S or sigmoid curve. To illustrate this
curve we present the following example using an algal reproduction graph (page 40) which has been adapted
from Wilson and Bossert (1971). Monthly algae cell densities and/or seasonal algal population growth (y
axis) in a tidal pool or creek are plotted over the course of a year (x axis). There are several steps to the
typical S growth curve. Between December and March when nutrients are limited and the temperatures are
low, there is little to no algae cell production. During the spring, warming temperatures and an increase in
nutrient loading causes the algal population to increase. This growth period is labeled “exponential growth”
or (r) in the graph. Typically, during the summer, there is an increase in nutrients including the nitrates and
phosphates from lawn fertilizers, golf courses, and farms (also known as nonpoint source pollution). The
abundance of these nutrients will cause algae to reproduce rapidly followed by a slowing trend and a
“leveling off” when the population approaches its carrying capacity (K). Carrying capacity occurs when
nutrients are at optimum levels and death equals algal production.
40 DRAFTAUGUST2015
#AlgalCells/m
l
ExponentialGrowth(r)
3,000,000
2,500,000
2,000,000
1,500,000
1,000,000
500,000
0
J F M A M J J A S O N D
Month
Figure Legend: Graph illustrating a Sigmoid (S) population growth curve by describing algae population growth (monthly
algal cell count per milliliter) over the course of a year in estuarine systems.
As depicted in the example above, the letters r and K represent the components of a population growth
curve derived by a formula. In order to understand population dynamics we must break apart the building
blocks of the population curve and plug in the r and K values. The logistic equation for the Sigmoid or S
population curve as defined by Wilson and Bossert (1971) is:
dN = rN(1-N) dt K
defined as: N = number of individuals
t = units of time
r = constant rate of population increase (births greater than deaths)
K = Carrying capacity of the environment.
CarryingCapacity(K)
NoGrowth
41 DRAFTAUGUST2015
Density-Dependent versus Density-Independent Factors
Other factors can enter the population equation in natural environments or in biological communities. The
growth of an animal population strives to reach stability or carrying capacity (K). However, in natural
systems populations encounter forces (known as density-dependent or density- independent factors) which
will affect their density and growth. Density-dependent factors are internal forces that operate within the
population. For example, infections, diseases, or stress related health problems can occur within an oyster
population. In another instance, too many individuals of a species will cause lack of space, and depleted
resources creating competition and/or stress within the population. These problems (density- dependent
forces) occur only when the density of a population reaches a critical level (Hickman et. al. 1984). Density-
independent factors occur outside of the population, examples include drastic changes that are
environmental in nature. For instance, extreme weather changes, unusually cold weather, hurricanes, or
drought conditions are examples of density independent forces acting against a population.
Comparison of r- and K-Selected Species
Biologists have categorized animals on adaptations they have developed to deal with density- independent
or density-dependent situations that arise and effect populations. K-selected species are animals or species
whose populations can survive controls that are density-dependent in nature. Conversely, r-selected species
are animal populations that have developed adaptations that are density-independent in nature (Hickman et.
al. 1984). Density-dependent or density-independent factors will affect all populations.
In the table below, notice the general characteristics of r-strategy species and K-strategy species.
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r-selected species K-selected species
Mature rapidly Mature slowly
Short-lived Long-lived
Many offspring Few offspring
Little to no care at birth Care for young at birth
Considered pests due to high densities May become threatened or endangered
Opportunistic Have stabilized populations
Few juveniles become adults Young usually reach maturity
Small size Large size
Examples: fiddler crabs, frogs Whales, birds
If we imagine a scale with r-species on one end and K-species on the other, many animals would fall on
either end of the scale. However, many animal species can show traits of both r-selected strategies and K-
selected strategies causing them to fall in between r or K on the scale (adapted from continuum concept in
Hickman et. al. 1984). In the example below, turtles and whelks are relatively long-lived, offer little care for
their many offspring, and few young reach maturity.
r-selected mixed r and K selected K-selected
mud snails sea turtles whales
skeleton shrimp whelks dolphins
fiddler crabs
Diversity in an Estuary
Estuaries are transition zones between fresh and salt water. The composition of plants and animals in the
estuary depends on various environmental conditions such as salinity, temperature, tidal fluctuations,
dissolved oxygen levels, turbidity, depth, substrate, and pollution. If we dissected and examined the diversity
of a salt marsh or a mangrove swamp, we would find various communities and habitats. Salinity, flooding,
and anaerobic soil conditions affect the plant communities within the estuary. In a salt marsh, smooth
43 DRAFTAUGUST2015
cordgrass (Spartina alterniflora) is most adapted to these harsh environmental conditions, so this species is the
most common. Areas of high marsh give rise to black needle rush (Juncus roemerianus). In mangrove swamps,
red mangroves (Rhizophora mangle) and black mangroves (Avicennia germinans) are the most adapted to these
harsh environmental conditions while white mangroves (Laguncularia racemosa) and buttonwoods (Conocarpus
erectus)are more common in higher portions of the estuary. In areas where water sits for a long period, and
evaporation rate is high, a layer of salt accumulates on the soil. This area (known as a salt pan) is where no
plants can grow due to very high soil salinities. In some zones of the high estuary, the salinities are too high
to support growth of smooth cordgrass or mangroves. Other plant species that can tolerate higher salinities
include glasswort (Salicornia spp.) and salt grass (Distichilis spicata).
Similar elevation zones within the estuary exist for animal communities. The higher elevations of the estuary
(near the estuary-forest edge) support shrubs and trees, providing cover for sparrows, marsh wrens, and
marsh hawks. Additionally, the higher elevations of the estuary are the location of the “wrack line” where
piles of dead vegetation accumulate and where spiders and amphipods reside.
The estuary’s bottom, called the “benthos,” supports a large community of invertebrates. Some inhabitants
include the mud snails (Nassarius obsoletus), fiddler crabs (Uca spp.), mud crabs (Rhithropanopeus harrisii), various
species of tanaids, isopods, oligochaetes, and polychaetes (worms)
such as Capitella capitata, Neanthes succinea and Streblospio benedicti.
Most of these species feed on sediment/detritus plus associated
bacteria and protozoans. Bivalves on the estuary floor filter feed on
plankton, algae, and bacteria. These bivalves include the ribbed
mussel (Geukensia demissa), Carolina marsh clam (Polymesoda
caroliniana) and eastern oysters (Crassostrea virginica). Marsh periwinkle
snails (Littorina littorina) graze on the blades of the salt marsh grass
(Spartina alterniflora).
Marsh periwinkles. Photo credit: Florida Sea Grant
44 DRAFTAUGUST2015
Changing environmental conditions can be stressful to aquatic organisms, and few species (but high
abundance of animals in each species) live in the estuary. However, species composition can fluctuate with
the seasons. Generally, peak densities of estuarine fauna occur in the spring or fall, with lowest animal
density occurring during the summer due to predation and competition. Typically, biodiversity is low in the
estuary and animal populations are usually r-selected species.
Diversity of Estuarine Mud Flats
Mud flats are another habitat supporting estuarine invertebrates and fish. They are located at the edges of
the salt marsh extending out into the creeks and rivers. These areas are comprised of varying combinations
of clays, silts, or sand. Although few species dwell in this zone, the densities of animals present in the mud
are high. Polychaetes and mud snails are generally the dominant organisms found in the mud. However,
there are various assortments of predators also found (usually at high tide). These include blue crabs
(Callinectes sapidus), grass shrimp (Palaemonetes pugio), penaeid shrimp (Penaeus aztecus, P. setiferus), silversides
(Menidia spp), and killifish (Fundulus spp). As the tide changes, many fish (e.g. skates, rays, and flat fish such as
flounders, small sharks, and red drum) will congregate on the shallow waters of a mud flat feeding on many
of the animals in the area.
Blue crab. Photo credit: Jarek Tuszynski/Wikimedia Commons
45 DRAFTAUGUST2015
Distribution and numbers vary according to the season, with the lowest numbers of organisms occurring
during the summer. This is possibly due to high predation as well as to high water temperatures and
associated low oxygen levels (Hackney et. al. 1992).
Diversity of an Oyster Reef
Oyster reefs in coastal Florida are common. However; they were even more prevalent in the early 19th and
20th centuries. Overfishing, mismanagement, disease, and poor water quality have taken a tremendous toll on
the oyster population. Oyster reefs located along tidal creeks and in estuarine rivers grow best between the
high and low tide line (intertidal). The common species is the eastern oyster/American oyster (Crassostrea
virginica). Oyster reefs can be quite a diverse community, supporting from 20 to up to 300 different species.
Oyster reefs and their associated species provide major food sources for numerous invertebrates and fish,
making oyster reefs a valuable habitat. For this reason oysters are termed “keystone species.”
Common species on an oyster reef include several species of mud crabs (Panopeus obesus, P. simpsoni, and
Eurypanopeus depressus) that feed on oysters and small crustaceans. Filter feeders such as the hooked mussel,
ribbed mussel, barnacles, and sponges attach to oyster shells. Several species of worms are present between
American oystercatcher. Photo credit: Alan D. Wilson/Wikimedia Commons/www.naturespicsonline.com
46 DRAFTAUGUST2015
shell crannies of the oyster reef, the most common worm being Neanthes succine. Other worms that are
present in this habitat include Polydora websteri, Heteromastus filiformis and Streblospio benedicti. Fish are often
found in oyster reef communities. Some of these include gobies (Gobiosoma spp.), blennies (Chasmodes spp,
Hypleurochilus spp., Hypsoblennius spp), skilletfish (Gobiesox strumosus), and toadfish (Opsanus spp). Predators
include whelks, flatworms, crabs, skates, rays, black drum, and the American oystercatcher (Hackney et. al.
1992). The oyster reef community and the associated species are major food sources making it a valuable
habitat to estuarine systems.
How is Diversity Measured?
A diversity index is often calculated to describe the diversity of animals present in a community. These
indices typically concern the measure of order or disorder within the ecosystem. The way it works is we ask
the question: How difficult would it be to predict the species of the next individual collected from the community? The
degree of uncertainty associated with this prediction is a measurement of diversity. If we feel confident in
naming the next species collected from a sample, the uncertainty number or diversity index is low (i.e. there
are so few species present that it would be relatively easy to predict the next species sampled). When the
diversity index value is high, the uncertainty value is high, making it more difficult to predict the next species
collected (i.e. there are so many different species present that the odds of guessing the next one collected are
very low). One of the simplest and most widely used diversity indices is the Shannon-Wiener Index (H’)
which takes species richness (number of species present) and species evenness (relative abundance of each
species present) into consideration. An example of how to calculate this index is provided on page 58.
The Shannon-Wiener index formula is:
H’= - ∑ Pi ln Pi OR = - sum of [(Pi)(Natural Log)(Pi)] for each species present i=1
Where Pi is the relative abundance (or proportion) of each species = ni/N ni = number of individuals in species i N = total number of individuals in all species
47 DRAFTAUGUST2015
Monitoring Protocol
1. Estuarine Bioassessment
When choosing a site, please select techniques from below if you would like you can do more than one.
A. D-Net Survey (every 3 months)
Based on methods described in: Biomonitoring and Management of North American Freshwater Wetlands
(Rader et al., 2001)
What you will need:
1. long rope and 2 PVC poles
2. yard stick or meter stick (1)
3. Adopt-A-Wetland Manual
4. buckets (2-3)
5. dishes or pans to sort the organisms
6. D-Net
i.) Set up the transect during a high tide in the high estuary or estuary border. A total of 5 survey
stations should be selected for each transect. Calculate your total distance from high to low estuary
and divide by 5 to determine the distance between survey stations. Remember to enter the estuary as
far as you can safely go.
ii.) Mark off a 1-meter section of the transect at each survey station. At each station hold the D-Net
parallel to the line of your transect. Sweep the D-Net along the sediment surface, scraping the mud
or sand and organic debris in the process. Perform sweeping motions 5 times along one side of the
1-meter section of the transect. Repeat the same procedure on the direct opposite side.
iii.) When you finish sweep netting at each station, sort through the debris, separate, count, and identify
the organisms. Combine all of the organisms from each survey station along the transect into one
sample. Record the information on biological survey worksheets (pages 53, 54, 55).
Checking a D-net. Photo credit: USFWS
48 DRAFTAUGUST2015
iv.) Groups can then calculate the diversity index for macroinvertebrates by completing the Biological
Diversity Index Worksheet on page 58.
v.) If possible, use a dock or tree that is easily identified as a starting point marked so that when
returning you can monitor roughly the same transect.
B. Quadrat Survey (every 3 months)
What you will need:
1. long rope and 2 PVC poles
2. one yard or meter quadrat (page 93 )
3. calculator
4. Adopt-A-Wetland Manual
5. buckets (2-3), and sorting pan
i.) Begin in the high marsh or marsh border. A total of 5 survey stations should be selected along a
transect. Station number one should be closest to the high estuary, followed by the other stations
extending out into the estuary. Calculate your total distance from high to low estuary and divide by 5
to determine the distance between survey stations. Remember to enter the estuary only as far as you
can safely go.
ii.) Set the quadrats along the transect at you determined survey stations. The quadrat will be the area
where you will work from so be careful not to step inside and disturb the area.
iii.) When at each station, sort through the debris, separate, count, and identify the organisms. Count all
organisms including clams, mussels, snails, crabs and crab holes that you find in the box. Include
organisms you find on the grass or vegetation, as long as they are in the quadrat. Identify and
combine all of the living organisms from each box and record the data on the Quadrat Survey Data
sheet, as it pertains to the station number (page 56). Groups can calculate the diversity index by
completing the Biological Diversity Index Worksheet on page 58.
Using a quadrat Photo credit: Maia McGuire
49 DRAFTAUGUST2015
iv.) Identify the different kinds of plants in each survey quadrat and count the total number of individual
plants of each kind and record on Quadrat Survey Data sheet (e.g. 20 Spartina grasses and 23 needle
rush) on page 56.
v.) Measure the height of 15 individual Spartina grasses and calculate the average for each survey station.
Record the height information on Cordgrass Height Data Sheet on page 57.
vi.) Quadrat surveys can be difficult in mangrove swamps we recommend surveying only swamp edges
with easy safe accessibility.
C. Hester-Dendy Survey (once per month)
Based on methods described in: Standard Methods Ed. (Eaton et al., 1995).
What you will need:
1. Hester-Dendy Colonizing Plates
2. buckets (2)
3. pliers
4. knife for scraping plates
5. rope from which to suspend plates
6. small dishes or pans to sort organisms
7. magnifying glass would be helpful
8. kitchen strainer
9. good lamp/lighting
10. Coastal Florida Adopt-A-Wetland Manual
Hester Dendy plates. Homemade (top) and commercial (bottom). Photo credit: Maia McGuire
50 DRAFTAUGUST2015
i.) Place the colonizing plates approximately six inches to one foot under the surface of the water
(measure from the top loop of the plates and the surface of the water).
ii.) Allow the plates to suspend in the water for one month. If left in the water too long, too many
organisms will colonize which makes it difficult to sort.
iii.) When the appropriate time has elapsed, retrieve the plates and take them to an area with proper
lighting so you can sort them according to groups.
iv.) Use the pliers to loosen the nut on the bottom of the samplers, pull all of the plates apart and scrape
both sides of each plate. Collect all of the debris into a bucket of salt water. If the water is too dirty,
sift the water and debris through a sieve. A metal kitchen strainer with a fine mesh works well.
v.) Sort all of the organisms that look alike, for example, all of the snails go in one dish/pan, crabs in
another. This process may take a while but you will improve your skills each time you collect.
vi.) By using the identification guide in the back of the manual, identify organisms and record on the
biological form on pages 54 and 55. Release any living organisms back into the river or tidal creek.
vii.) Advanced groups can calculate the diversity index on page 58 for macroinvertebrates by completing
the Biological Diversity Index Worksheet in the manual.
viii.) To re-assemble the Hester-Dendy colonizing plates: you should have 24 spacers and 14 disks;
starting from the “eye loop” (top) put 9 disks separated by 1 spacer each, separate disk 10 by 2
spacers, 11-12 disks by 3 spacers, and 13-14 by 4 spacers. When fully assembled secure with the
washer and wing nut.
2. Beach Bioassessment
Please select techniques from below. If you would like, you can do more than one.
A. Hester-Dendy Survey
If there is a dock piling or similar structure you can use to suspend the Hester-Dendy plate sampler from,
use this technique by following the instructions provided above.
51 DRAFTAUGUST2015
B. Seine Survey (every 3 months)
Seining is a commonly-practiced technique wherein a net is drawn through the water to capture the
organisms.
What you will need:
1. seine
2. buckets (3)
3. Adopt-A-Wetland manual
4. yard stick (to measure depth)
i.) At the beach when approaching the surf, always stay at a safe depth - preferably 3 feet. Sweep
through the shallow water with a 10-20-foot seine for approximately 2-3 minutes. Repeat this
procedure 3 times.
ii.) Identify fish and invertebrates by using the identification guide and record your results in the
worksheets on pages 54 and 55.
iii.) Groups can calculate the diversity index by using the Diversity Worksheet (page 58).
3. Mangrove Diameter and Height Measurements (CARICOMP Methods Manual, Levels 1 & 2 March 2001)
What you will need:
1. 1 m long cloth tape measure�
2. 6 m telescopic measuring rod
3. Pencil/Data Sheet
Checking the contents of a seine. Photo credit: GTMNERR
52 DRAFTAUGUST2015
Diameter - Measure the circumference of the tree in order to obtain diameter of the trunk (which is a
standard measure used by foresters; normally expressed as diameter at breast height or dbh. Breast height is
considered to be 4.5 feet). With red mangroves, the circumference (c) is measured immediately above the
buttress roots, using a flexible tape marked in centimeters. Diameter is then calculated as:
dbh = c/п
Red mangrove trees sometimes have more than one trunk arising from a common buttress or "prop-roots."
In these cases, each trunk is measured as a separate tree. Ignore prop-roots growing down from high
branches when deciding where to measure the circumference.
Height - should be measured for all trees in the plot using three parameters (a) height above sediment
surface of the highest prop root, (b) length of trunk, from prop roots to main area of branching and (c) total
height, from ground to highest leaves. For saplings and trees up to 6m, a graduated telescoping rod is used.
Where tree density is high, measuring height may be very difficult. Estimate as closely as possible, where it
may be difficult to obtain actual measurement. You can use a technique involving geometry if you want to
be fairly accurate about your height estimation.6
6http://forestry.usu.edu/htm/kids-and-teachers/tree-height-measurement
53 DRAFTAUGUST2015
COASTAL FLORIDA ADOPT-A-WETLAND BIOLOGICAL MONITORING FORM
AAW Group Name County
Group ID- Site ID
Investigators
Wetland Name Date Time
Rain in Last 24 Hours? Yes/No Amount of Rain Inches in Last hours/days
□Heavy Rain □Steady Rain □Intermittent Rain
Present Conditions: □Heavy Rain □Steady Rain □Intermittent Rain
□Partly Cloudy □Overcast □Clear/Sunny
Site Location Description (e.g. Salt Marsh, Mangroves, Beach)
Sampling Technique: □D-Net □Quadrat Survey
□Seine □Hester Dendy (How long in water?)
Notes:
54 DRAFTAUGUST2015
GROUP NAME: SITE NAME: DATE:
Coastal Adopt-A-Wetland Biological Community Sampling Form
Phylum Mollusca Phylum Arthropoda Phylum Ecinodermata Class Gastropoda (Snails & Slugs) Class Cirrepedia (Barnacles) Class Holothuroidea (Sea Cucumbers) Oyster Drill Barnacle Sea Cucumber Mud Snail Class Malacostraca (Crabs, Shrimp) Class Asteroidea (Sea Stars) Knobbed Whelk Fiddler Crab (sand, mud, brackish sp.) Sea Star Lightning Whelk Mud Crab Class Echinoidea (Sea Urchins, Sand Channeled Whelk Blue Crab Key Hole Urchin (Sand Dollar) Tulip Snail Hermit Crab Sea Urchin Dove Snail Stone Crab Class Ophiuroidea (Brittle Stars) Rock Snail Porcelain Crab Brittle Star Keyhole Limpet Spider Crab Phylum Annelida Nudibranch Calico Crab Class Polychaeta (Worms) Lettered Olive Speckeled Crab Worm Class Bivalvia (Mussels, Clams, Oysters) Class Merostomata (Horseshoe crabs) Class Hirudinea (Leeches) Ribbed Mussel Horseshoe Crab Leech Hooked Mussel Class Pycnogonida (Sea Spiders) Scorched Mussel Sea Spider Paper Mussel Hard Clam Phylum Cnidaria Surf Clam Class Anthozoa (Anemones) Oyster Anemone Ark Sea Whip Jacknife clam Sea Pansy Coquina Other Marsh clam Class Scyphozoa (Jellyfish) Dwarf Surf clam Jellyfish Phylum Porifera Other Class Demospongiae (Sponges) Redbeard Sponge Phylum Ctenophora Basket Sponge Class Tentaculata (Comb Jellies) Finger Sponge Comb Jellies Total Number of All Kinds: Boring Sponge Other Total Number of All Individuals:
55 DRAFTAUGUST2015
GROUP NAME: SITE NAME: DATE:
Coastal Adopt-A-Wetland Biological Community Sampling Form
Phylum Chordata Lined Seahorse Spanish Mackerel Class Ascidiacea (Tunicates, Sea Squirts) Lookdown Spot Sea Squirt Mosquitofish Spotted Hake Sea Grape Mummichog Spotted Seatrout Sea Pork Naked Goby Star Drum Other Northern Needlefish Striped Anchovy Class Osteichthyes (Bony Fishes) Northern Pipefish Striped Blenny American Eel Northern Puffer Striped Burrfish Atlantic Bumper Northern Sea Robin Striped Killifish Atlantic Croaker Ocellated Flounder Striped Mullet Atlantic Cutlass fish Oyster Toadfish Striped Sea Robin Atlantic Menhaden Pigfish Summer Flounder Atlantic Silverside Pinfish Tarpon Atlantic Spadefish Planeheaded Filefish Weakfish Atlantic Thread Herring Red Drum White Mullet Bay Anchovy Rock Sea Bass Whiting Big Head Sea Robin Sailfin Molly Windowpane Black Drum Sand Perch Other Black Sea Bass Sea Catfish Blackcheek Tonguefish Sharksucker Class Elasmobranchiomorphi Bluefish Sheepshead Atlantic Sharp Nose Shark Butterfish Sheepshead Killifish Atlantic Stingray Crevalle Jack Silver Jenny (Mojarra) Bonnet Head Shark Feather Blenny Silver Perch Clearnose Skate Florida Pompano Silver Seatrout Lemon Shark Gafftopsail Catfish Skillet Fish Sandbar Shark Goby Smooth Puffer Smooth butterfly Ray Gray Snapper Southern Flounder Southern Stingray Hogchoker Southern Harvestfish Other Inshore Lizardfish Southern Sennet Total Number of All Kinds: Ladyfish Southern Stargazer Total Number of All Individuals:
56 DRAFTAUGUST2015
Quadrat Survey Data Date: AAW Group Name: Length of Transect (ft or m): Identify the types and numbers of animals and plants in each survey station (for help in identification please see Macroinvertebrate and Plant Identification keys in the Appendix of this manual).
Station 1 # Station 2 # Station 3 # Station 4 # Station 5 # Atlantic Ribbed Mussel
Atlantic Ribbed Mussel
Atlantic Ribbed Mussel
Atlantic Ribbed Mussel
Atlantic Ribbed Mussel
Eastern Oyster Eastern Oyster Eastern Oyster Eastern Oyster Eastern Oyster
Periwinkle Snail Periwinkle Snail Periwinkle Snail Periwinkle Snail Periwinkle Snail
Mud Snail Mud Snail Mud Snail Mud Snail Mud Snail
Coffeebean Snail Coffeebean Snail Coffeebean Snail Coffeebean Snail Coffeebean Snail
Amphipod Amphipod Amphipod Amphipod Amphipod
Fiddler Crabs Fiddler Crabs Fiddler Crabs Fiddler Crabs Fiddler Crabs
Crab Holes (> ¼ inch)
Crab Holes (> ¼ inch)
Crab Holes (> ¼ inch)
Crab Holes (> ¼ inch)
Crab Holes (> ¼ inch)
Cordgrass (Spartina)
Cordgrass (Spartina)
Cordgrass (Spartina)
Cordgrass (Spartina)
Cordgrass (Spartina)
Needlerush (Juncus)
Needlerush (Juncus)
Needlerush (Juncus)
Needlerush (Juncus)
Needlerush (Juncus)
Seaweed Seaweed Seaweed Seaweed Seaweed
☺Helpful hint: If there are too many grasses in the box you can estimate the number of grasses.
57 DRAFTAUGUST2015
Cordgrass (Spartina) Height Data Sheet
Date: AAW Group Name:
Measure height of 15 Spartina plants in each survey station and choose color (green, yellow, brown) for each stem measured then calculate the average.
Spartina Sample #
Station #1 Station #2 Station #3 Station #4 Station #5 Ht Color Ht Color Ht Color Ht Color Ht Color
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 Sum Average
Height
Optional
Please estimate the number of blades of Spartina in your stations and circle the appropriate color option.
Station 1
Station 2
Station 3
Station 4
Station 5
< 10 Green < 10 Green < 10 Green < 10 Green < 10 Green
20-50 Yellow 20-50 Yellow 20-50 Yellow 20-50 Yellow 20-50 Yellow
> 50 Brown > 50 Brown > 50 Brown > 50 Brown > 50 Brown
58 DRAFTAUGUST2015
Shannon-Wiener Biological Diversity Index (H’) Worksheet
H’= - S∑ Pi ln Pi OR = - sum of [(Pi)(Natural Log)(Pi)] for each species present i=1
Where Pi is the relative abundance (proportion) of each species = ni/N ni = number of individuals in species i
N = total number of individuals in all species S = number of species
1. Circle type of monitoring: Box Survey, Colonizing plates, D-Net, and Seine 2. Habitat type (i.e. oyster reef, salt marsh): 3. Calculate the diversity index for your sample by completing the worksheet below.
A B C D E F Species
(i)
# Individuals
of Each Species
(ni)
Total Number of
Individuals in all Species
(N)
Relative Abundance
Of Each Species
(Pi)
Natural log of
Relative Abundances
(ln Pi)
Relative Abundances
Times Their Natural log
(Pi ln Pi)
e.g. Mud Crab 2 39 0.05 -2.99 -0.15
Sum of Column F = Multiply by -1 to make positive = Shannon-Wiener Index Diversity Index =
Diversity Scale (circle appropriate value for your site)
1 2 3 4 5
Not Diverse → Moderately Diverse → Diverse
59 DRAFTAUGUST2015
Mangrove Diameter and Height Measurements
AAWGroupName:
Date:
Location/GPS:
MangroveSpecies(red/black/white)
Circumference(c) Diameter(c/pi)
Highestproproot Firstbranches totalheight
Comments:
60 DRAFTAUGUST2015
Physical/Chemical Parameters
Physical/Chemical Monitoring Protocols
Physical/Chemical Survey Worksheets
ChapterFivePhysical/ChemicalMonitoring
Photo credit: Helle Patterson
61 DRAFTAUGUST2015
Physical/Chemical monitoring is conducted at regular intervals at the same location. This level of
monitoring can be used to gather information about specific water quality characteristics. Regular
monitoring helps assure that your information can be compared with changes over time. In tidally-
influenced locations be sure to note the direction of the tide (incoming/outgoing) and conduct sampling at
the same tidal stage each month. At extremely low tides there may not be water present for sampling, so
check tide charts before going to your site. Also, chemical testing during or immediately after a rain may
produce very different results than during dry conditions. Therefore, it is very important to record weather
conditions. If conditions are unsafe for any reason, DO NOT SAMPLE.
Several physical/chemical conditions may be tested at your site including temperature, pH, dissolved oxygen
(DO), salinity, turbidity, and settleable solids. Be sure to follow all instructions and safety guidelines when
conducting chemical analysis.
Physical/Chemical Parameters
Temperature
Temperature has an effect on the chemical and biological processes of an aquatic system. Temperature will
affect the dissolved oxygen level, density of water, as well as the distribution of organisms and their
metabolic processes. Temperature differences between surface and bottom waters can produce vertical
currents, which will transport or mix nutrients and oxygen throughout the water column.
Changes in temperature also occur with depth. These changes result from cooling and warming
temperatures during the seasons. In the warm summer months, warmer water is on the surface with cooler
water at lower depths. When air temperatures begin to cool (as in late autumn), the surface water becomes
Physical/ChemicalMonitoring
62 DRAFTAUGUST2015
cool and dense and sinks to the bottom. This mixing of the surface and bottom layers of water will cause
nutrientsto disperse from the bottom to the surface. The dispersed surface nutrients act as fertilizers for
phytoplankton. Phytoplankton are free-floating microscopic aquatic plants drifting in surface areas of the
water. Phytoplankton blooms usually occur in warmer temperatures during the spring, summer, or fall
(Ohrel and Register, 1993).
Water temperature is also influenced by wind, storms, and currents created by tides. The water movement
created by currents or wind causes a higher rate of mixing within the entire water column. Tributaries such
as rivers or shallow tidal creeks, which respond quickly to atmospheric temperature, may influence
temperature as they flow into the estuary or tidal creek. Also, water temperature may be increased by
discharges of water used for cooling purposes or by runoff from heated surfaces such as roads, roofs, and
parking lots. Also, cold underground water sources (i.e. springs) and the shade provided by overhanging
vegetation could lower water temperatures in some areas.
pH
We measure the pH of water to determine if it is acidic or basic (alkaline). Chemically speaking, the “p” in
pH refers to the “potential” of the H+ ion. There can be a high concentration of H+ ions or a high
concentration of OH- ions. The higher concentration of H+ ions causes the sample to have a lower pH
(acidic). The higher concentration of OH- ions in a sample will cause the pH value to be basic (above 7)
(Wetzel and Likens, 2000). The pH measurements are recorded on a scale from 0 to 14, with 7.0 considered
neutral. Solutions with pH below 7.0 are considered acidic; those between 7.0 and 14.0 are considered basic.
The pH scale is logarithmic, so every one-unit change in pH represents a ten-fold change in acidity. In other
words, pH 6 is ten times more acidic than pH 7; pH 5 is one hundred times more acidic than pH 7.
63 DRAFTAUGUST2015
ACIDIC
BASIC
pH Example
0.5 Battery Acid
2.0 Lemon Juice
5.9 Rainwater
7.0 Distilled Water NEUTRAL
8.0 Salt Water
11.2 Ammonia
12.9 Bleach
Chemical/physical characteristics of the water or substrate can influence the pH. Monitoring the pH of
water gives us an indication of the health of our estuaries. As a thermometer is to human health, telling us
when we are sick, the pH tells us if something is unhealthy about our aquatic systems. An abnormal pH
reading indicates that the system is chemically out of balance. However, the pH range of any coastal wetland
may be highly variable depending on several factors such as rainfall, plant/bacteria growth, temperature, or
salinity. The salinity of the water affects the pH in our coastal estuaries. The pH and salinity follow a pattern
from fresh water river input to offshore areas. Coastal areas with freshwater influence (low salinity zones)
will have a pH range of 7.0 to 7.5 and in areas of higher salinities (offshore), pH ranges between 8.0 and 8.6
(Ohrel and Register, 1993).
The buffering capacity of carbonates and bicarbonates in seawater will increase pH values. Biological activity
can suddenly alter the water chemistry causing increases or decreases in pH values. For example, rapidly
growing algae will produce oxygen and remove carbon dioxide (CO2) from the water during photosynthesis
resulting in an increase of pH. Conversely, decomposition of organic matter or respiration in plants and
animals will use up oxygen but increase CO2 levels in the water thus, lowering the pH values. Abnormally
low or high pH values can adversely affect egg hatching and larval development, stress fish and insects, and
even cause fish kills (Meadows and Campbell, 1978). Some human factors influencing pH readings outside
the normal range include mine drainage sites, atmospheric deposition or industrial point discharges. Serious
64 DRAFTAUGUST2015
problems occur in coastal waterways when the pH falls below five or increases above 9 (Ohrel and Register,
1993).
Soil and Sediment pH
Estuarine environments produce large amounts of dead vegetation or organic matter especially during the
winter when leaf dieback is common. The organic matter lies on the estuarine floor decomposing. During
decomposition, dead grass and leaves are broken down by bacteria that give off carbon dioxide (CO2)
during respiration. The increased CO2 production lowers the pH of the sediment. Generally, the pH of
sediment in estuaries, tidal creeks, mangrove swamps, and salt marshes reflect that of the water above. In
salt water, the pH can be 7 to 8.5, and is generally the same in the sediments. However, there are slight
fluctuations daily due to processes such as photosynthesis and respiration. Throughout the day, algae on the
sediments photosynthesize, increasing oxygen production, which increases the pH of the soil. During the
night, respiration increases carbon dioxide which shifts the pH lower. Sediment pH can fluctuate from 9 or
10 during the day time to 7 or 8 during late evenings. Although the pH fluctuations vary according to algae
densities, shade from marsh grass, and season, the shifts may be more noticeable according to conditions of
the area (Pomeroy, 1959).
Dissolved Oxygen
Dissolved oxygen (DO) is the most critical factor in determining the health of an aquatic system. Dissolved
oxygen is the measurement of the oxygen content in the water. Sources of oxygen in aquatic systems include
atmospheric diffusion, plant/algae photosynthesis, currents, and wave action. Oxygen is required for
respiration in animals and plants. Oxygen is also consumed during decomposition of organic matter by
respiring bacteria.
Shifts in DO are related to the time of day, season, and temperature. There is a higher oxygen production
during the day when algae is producing oxygen through photosynthesis. During the night when there is no
65 DRAFTAUGUST2015
light source, photosynthesis stops while respiration continues, thus shifting oxygen levels lower than during
the day.
Another factor that influences oxygen levels is salinity. Water with high salinity holds less oxygen than water
with lower salinity levels. In addition, temperature affects dissolved oxygen levels. Cooler water contains
more oxygen than warmer water for any given salinity. For instance, in the cooler months the dissolved
oxygen levels can increase to 8 or 9 parts per million (ppm). Conversely, the warmer waters during the
summer months cannot hold oxygen as readily so levels decrease to 4 ppm and in some cases as low as 3
ppm. An ecosystem with low oxygen levels can be further stressed when excessive nutrients from fertilizer
runoff are added into the tidal creek or estuary. Nutrients can trigger an algal bloom which will increase
oxygen temporarily, but when the algae die problems may occur. Respiration and decomposition during the
algae die-off will further deplete oxygen levels causing stress on aquatic life. In general, low oxygen levels
indicate a stressed aquatic environment. When a system is further stressed by an input of pollutants, sewage,
or organic matter, decomposition of these materials will seriously deplete oxygen levels (Ohrel and Register,
1993).
Low oxygen levels are occurring in a “Dead Zone” at the mouth of the Mississippi River in the Gulf of
Mexico and increasing in occurrence along the eastern seaboard. These low oxygen levels worsen in the
summer. High temperatures reduce the amount of oxygen that the water can hold, causing the dead zone to
increase in size. The dead zone is possibly a result of industries, farms, and residential areas that collectively
use fertilizers. Fertilizers are carried by inland streams, wash into water ways, and are eventually
concentrated and transported to the coast. As the concentration of certain fertilizers/nutrients increase on
the coast, it causes an algae bloom. A bloom is a large concentration of microscopic algae (enough that it
will discolor coastal waters and limit light penetration for plants living on the bottom). The bloom becomes
so dense that the algae deplete the nutrients and the population crashes. The algal population crash causes a
sudden burst of decomposition, which involves bacteria breaking down algal cells. The problem arises at
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this point as oxygen is used in this process causing severe oxygen depletion. Dissolved oxygen < 2 ppm is
termed hypoxia, which is lethal to most fish and causes bivalves to use anaerobic respiration; dissolved
oxygen < 0.5 ppm is termed anoxia ̶ devoid of oxygen.
Salinity
Everyone wonders why the ocean water is salty. Imagine the ocean as a “sink”, where material carried by the
creeks and rivers collects. Over time and seasons, through heavy rains and wind, the rocks and mountains
slowly wear away. Many of these small particles or minerals come from chunks of mountains and rock. Over
time, they are transported via the waterways and are deposited in the ocean or “sink”. So, most of this
material dissolved in the water had its origin further inland. Looking closely at the transported materials
present in the ocean, there is a combination of minerals, elements, and salts. Examples of these elements
and/or dissolved salts are chloride, sodium, sulfate, magnesium, calcium, and potassium. These dissolved
salts in the water make up the total salinity concentration. Approximately 85% of the salts in seawater come
from a combination of sodium and chloride (Na is 30.6%) and (Cl 55%) which makes up table salt ― NaCl.
The remainder of dissolved salt constituents include magnesium (Mg 3.7%), sulfate (S04 7.7%), potassium (K
1.1%), calcium (Ca 1.2%) and silica (Si) (Coulombe, 1992). Organisms such as those with shells, for example
clams and snails (phylum Mollusca) and sea urchins (phylum Echinodermata), absorb calcium from the
seawater to incorporate and build their strong shells. Microscopic algae or diatoms absorb silica from the
seawater to build their glasslike floating bodies (Greene, 2004). The total salts or salinity is measured in parts
per thousand (ppt, ‰). Seawater has an overall average salinity of 35 ppt, or every 1000 pounds of water
contains 35 pounds of salt (Thurman, 1987).
The salinity concentration in estuaries increases as you go toward the ocean. You can think of salinity
concentration in zones. A freshwater river will enter the coastal area and may have little to no salinity
(ranging from 0.5 to 5 ppt). The freshwater then mixes with salt water and becomes part of the estuary. An
estuary is defined as a place where salt and fresh water mix. Salinities in an estuary will vary from 5 to 18 ppt
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closest to the freshwater input, to 18 to 30 ppt or higher when closer to the open ocean (Ohrel and Register,
1993).
Salinities fluctuate with weather conditions, tides, and river flow. Salt water is a somewhat hostile
environment for animals and plants to inhabit. High salinities often cause diseases in oysters (O’Beirn et al.,
1994). Some organisms can adapt well to fluctuating salinities. They might move to other areas, burrow into
the sediment, alter their osmotic pressure by producing more or less urine, or drinking more or less fresh
water. Some organisms that can adapt to changing salinities include blue crabs, oysters, shrimp, mussels, and
mullet. Salt marsh plants must endure high salinities in the marsh soils, so the few species that live in the
marsh have certain adaptations. For instance, one species that dominates the salt marsh is the smooth
cordgrass or Spartina alterniflora. Smooth cordgrass has an adaptation (similar to desert plants) in that it can
conserve water by adjusting its osmotic vascular pressure. This is done by maintaining a high concentrate of
other organic solutes found in marsh soils and by discharging excess salts through tiny salt glands or pores
located on the blades of the leaves. In fact, if you visit a salt marsh and closely observe the smooth
cordgrass, you can see the salt crystals on the leaf blade excreted by the salt glands. Mangroves are also
highly adapted to life in saline water and soil; their salts are either excluded or excreted by the plant.
A refractometer measures the dissolved solids or salts in seawater. Salinity in water can be observed by how
light interacts with a sample when a drop is placed on a prism. Refraction is the change in direction of a
light pathway when passing through two media (i.e. air and water) with different densities (Thurman, 1987).
A sample with high salinity has high refraction properties or “light bending abilities.” When peering through
the eye piece there are two scales that measures salinity 1) parts per thousand (ppt) located on the right in
most refractometers and 2) specific gravity (density) which is on the left side of the field of view. We use the
ppt scale when determining the salinity of a sample.
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Settleable Solids
Settleable solids include all suspended particles in the water. These particles naturally occur in coastal waters,
but at times an overload of sediment/particles will cause problems in estuaries and tidal creeks. Examples of
solids entering a river or creek include sewage or soils eroded from development on tidal creeks or rivers.
Excess solids in water block sunlight throughout the water column compromising the growth of aquatic
plants living on the bottom. These sediments can also clog the gills of fish and macroinvertebrates, limiting
their ability to extract oxygen from the water. Sediment can carry harmful substances such as bacteria,
metals, fertilizers, and pesticides from garden runoff. Settleable solids and turbidity tests give indications of
sediment load in the water (Ohrel and Register, 1993). A settleable solids test is a quantitative method to
determine if there is an overload of sediment or other solids in the water. The settleable solids test is not the
same as turbidity because a settleable solids test only measures larger, heavier particles that settle out in an
Imhoff cone after a 45 minute time period.
Turbidity
Turbidity is a measurement of water clarity (clearness). Seasons,
weather conditions, algal blooms, and the amount of suspended
particles in the water can affect turbidity. Turbidity includes all
particles in a sample. Phytoplankton, detritus, silt/clay particles, and
other organic matter contribute to the turbidity of water. The
higher the turbidity in the water, the less light penetrates into the
lower depths causing a decrease in growth of phytoplankton and
aquatic plants that are growing on the bottom. As phytoplankton
densities decrease in the water column, the rate of photosynthesis will slow causing oxygen levels to
decrease (Ohrel and Register, 1993). The “secchi disk” is used to measure water turbidity in the field (see
page 72)
Secchi disk: Photo credit: Mark Hoyer
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Physical/Chemical Monitoring Protocol
Temperature
Equipment: Thermometer/ water sampling bottle or bucket (optional)
Procedure:
1. Record the air temperature first by placing the thermometer in a shady area.
2. Water temperature may be taken from the water directly or from a large sample container (if done
quickly so the temperature does not change).
3. Record the information on the physical/chemical survey water monitoring form (page 78).
pH
Equipment: pH pen (sometimes will break down after using for 1 year). Please call if the pH pen you are
using is not working properly. The pH pens provided can also give the water temperature.
Procedure: To test pH in your water sample you follow these simple directions. For more information refer
to the directions that comes with the instrument or contact your Adopt-A-Wetland coordinator.
Calibrating the pH Pen (also see instructions included with the pen)
The pH pen needs to be calibrated monthly. We only use the one point calibration (only use the 7.0 buffer)
for salt water. To calibrate, press and hold the on/mode button until OFF on the lower display is replaced
by CAL. Immediately release the button. The display enters the calibration mode displaying “pH 7.01
USE.” Dip the pH pen in the 7.0 pH buffer. After 1 second, the meter responds, if the correct buffer is
detected then its value is shown on the display (pH 7.0 or 7.01) and REC appears in the lower part of the
display. There is a clock illustration on the upper left of the screen. Wait until the clock disappears then
press the on/mode button, at this point, the buffer is accepted. After the calibration point has been
accepted, the on/mode button must be pressed in order to return to the normal mode.
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1. To get a pH reading from your sample, remove cap from the electrode, press “On/Mode” button to
turn on the pH pen the display screen will be visible followed by a percent indication of the remaining
battery life (E.G. % 100 Batt).
2. Dip the electrode into the sample while stirring it slowly. Measurement is taken when the stability
symbol (looks like a clock) in the top left corner of the display screen disappears.
3. Note the pH, or you can press “Hold/Con” to freeze the reading. Record the pH value on your data
sheet.
4. Press “On/Mode” to turn the tester off.
5. Rinse electrode with tap water, but do not dry. Keep a damp sponge in the cap moist with the storage
solution.
Dissolved Oxygen
Equipment: Dissolved Oxygen test kit
Procedure: Also see instructions enclosed with LaMotte kit.
1. Submerse entire water collection bottle until all bubbles are out of the bottle, while bottle is still
submersed, cap it and tighten lid.
2. When adding solutions to water sample, do not touch the tip of the chemical bottle to the water sample.
3. Uncap bottle cap of the water sample and add 8 drops of Manganous Sulfate and 8 drops of Alkaline
Potassium Iodide.
4. Cap bottle and invert several times then do not disturb sample. Allow the precipitate to settle to the
bottom of the bottle.
5. When the brown precipitate has settled past the shoulder of the bottle, you are ready for the next step.
This may take up to 20 minutes.
6. Add 8 drops of Sulfuric Acid; be extra careful with this solution.
7. Cap and gently invert the bottle. The color will be an amber color, like apple juice.
8. Invert bottle until there are few to no brown particles remaining.
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9. Pour mixture from bottle into the titrating tube, up to 20 milliliters then add 8 drops of starch indicator
solution.
10. Color will turn dark blue. Fill syringe up to the 0 mark with Sodium Thiosulfate, make sure you
eliminate all bubbles.
11. Slowly add 1 drop at a time and gently swirl sample; color should turn from dark blue to clear. When
sample turns clear note the level of liquid left in your syringe as your dissolved oxygen amount (ppm) in
your sample. Note: Do not add too much Sodium Thiosulfate. Discard any leftover chemicals in the
waste bottle.
12. Record the information on the physical/chemical survey water monitoring form (page 78).
Water Salinity
Equipment: Refractometer
Procedure:
1. Lift lid on refractometer that protects the glass prism.
2. Place one or 2 large drops of sample onto the glass and close the lid.
3. Look through the eye piece and focus your view of the scale inside. Read the line where the blue color
meets the white color in the field of view.
4. Read the scale on the right-hand side that shows parts per thousand or ppt , ‰.
5. Rinse glass with tap water, blot dry with a paper towel, and store in case.
6. Record the information on the physical/chemical survey water monitoring form (page 78).
7. The refractometer periodically needs to be calibrated. For questions or help contact your local Adopt-A-
Wetland coordinator.
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Soil Salinity
Equipment: Refractometer
Procedure:
1. Dig a hole to a depth of 15 cm deep and allow the hole to fill with water. Note: You can use a pvc pipe
marked off at 15 cm to get water from this depth also.
2. As the water fills the hole/pvc pipe, use a small dipper or cup to retrieve a water sample.
3. Allow sediment to settle to the bottom of the cup.
4. Use a dropper to obtain a small amount of water and place water drops on the glass surface of the
refractometer.
5. Follow the directions for using the refractometer.
6. Record information on physical/chemical survey water monitoring form (page 78).
Settleable Solids
Equipment: Imhoff cone & stand
Procedure:
1. Pour one liter of sample water into cone.
2. Let cone stand at least 45 minutes.
3. Record level of solids.
Turbidity
Equipment: Secchi disk
Procedure:
1. Calibrate your line by first attaching the line to the Secchi disk. Mark the line every 10 cm by using a
permanent marker and measuring tape. While standing at your site (bridge, boat, or dock) lower the
Secchi disk into the water until it disappears. If possible, look for a shady spot to increase visibility. It is
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important that the disk travels vertically through the water column and is not “swung out” by the
current. Attach weights if necessary to prevent current pull.
2. Raise the disk slowly up and down several times until you find the point that the disk vanishes. Mark the
spot on the line where it enters into the water.
3. Calculate the depth at which the disk disappeared. This measurement is referred to as Secchi disk depth.
If disk goes all the way to bottom of water before disappearing the Secchi depth is greater than the
water depth and should be noted on the form.
4. Record the information on the physical/chemical survey water monitoring form (page 78).
Physical Monitoring Protocol for Beach Sites
Beach Slope Measurement: Emery Method for Beach and Dune Profiling7
Equipment: Transect Tape, Profile Stake, Emery Rod, GPS, Compass, Pencils, Clipboard, Data Sheet,
Camera
Procedure:
1. Ideally, plan to go at low tide during a full or new moon. Plan to measure to the water’s edge
(average out waves and run-up) or beyond if safe.
2. Features to note:
a. Breaks in slope;
b. Edge of dune vegetation (if any);
c. Fence position;
d. LHTS, last high tide swash mark;
e. WL, water line and
f. TIME measured.
7Emery, K.O., 1961, A simple method of measuring beach profiles. Limnology and Oceanography, v. 6: 90-93.
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3. Keep horizontal intervals regular. If using meters and a measurement is less than 1 m, then make the
next measurement a distance to return to a meter increment.
4. Walk next to the profile, not on it. This is particularly important for the lead person.
5. Write the TIME of last measurement in the Notes column when done. Complete the sketch.
6. Take photographs along the beach (shore-parallel direction) and one looking up the profile. It helps
to place the profile rods on the profile line and include them in the photograph.
7. Find the Starting Point. Set a control point (a reference stake or pin) in the ground. This is done
once before the first profile is taken. The same control point is reused for each subsequent profile and is the starting
point of all measurements. Take a compass reading from the control point perpendicular to the water’s
edge.
8. Run the Transect Line. Have one person hold the end of the transect line (measuring tape) at the
starting point. Another person should run the transect line following the desired compass reading so
the line is set perpendicular to the water’s edge.
9. Begin Notes. Fill in the top part of the log sheet. Include names of people in the team, the date,
time, profile name or number, beach location, etc
10. Record Stake Height. Measure the height of the ground in relation to the top of the control point
with the numbers (scale) up.
11. Set Rod 1. Stand the end of one profile rod (Rod 1) on the ground next to the control point with
the numbers (scale) up. The person holding this rod should stand off the profile line for the next
step.
12. Set Rod 2. The second person takes Rod 2 toward the ocean. Looking back toward land and Rod
1, this lead person places Rod 2 (with scale up) on the transect line. Pick a horizontal distance of a
meter (or other suitable distance if obstacles are in the way) as a spacing between the two poles. Use
a graduated chain or pole to do this and be careful to hold both poles straight up and down while
setting Rod 2 in place.
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13. Measure and Record. From the landward pole, the first person sights the horizon and the top of
the lower of the two rods. This line-of-sight will intersect part way up the other rod. Read the
elevation number marked on the other rod that is in line with the pole top and the horizon. Keep both
poles vertical when reading! Note that sometimes the reading will come from Rod 1 and sometimes from
Rod 2. This is because the ground may slope down or up and may change which pole is higher at
different places on the beach profile line. When the ground slopes down toward the ocean, the
forward rod (Rod 2) will be lower, and a negative [-] number is assigned to the vertical reading off of
Rod 1. When the ground slopes up looking toward the ocean, the forward rod will be higher, and a
positive [+] number is assigned to the reading. In this case, the number is read off the forward rod
(Rod 2). So moving forward on the profile, uphill is [+] and downhill is [-]. Always use either a + or
– before the number. It takes careful attention to get this right on each measurement. A single error
will make the rest of the data plot incorrectly on a graph. Record the elevation change and
horizontal distance between poles on the log sheet. Also note any features at the forward rod (such
as edge of dune, slope change, water line, etc.) in the Notes column on the log sheet.
14. Move Ahead. After the notes are taken, move Rod 1 to the same “footprint” occupied by Rod 2.
The person at Rod 2 should wait for Rod 1 to come up alongside Rode 2 in order to be certain of
getting the position correct. After Rod 1 is in the place of Rod 2, the forward rod can be moved
ahead another meter or two and placed on the ground in line with Rod 1 and the original control
point(s).
15. Repeat Steps 6 and 7. Measure, Record, & Move. Continue to move ahead, repeat these steps
all the way to the water. As you go, everyone on the team should look ahead for features to stop on
and measure. If some feature, perhaps the edge of the dune, does not occur at a horizontal interval
of one meter, then make the horizontal distance smaller. For example, if the dune edge is only 0.6m
from the least measurement, move the forward pole ahead only that far. ON the next measurement
move ahead only 0.4 (or 1.4) m in order to get back on a spacing of 1 m intervals. Keeping a set
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interval in whole meters will help with data analysis later.
16. Stop at the Water. Make a measurement that includes the water line. IN the notes column of the
log sheet abbreviate it W.L. and record the TIME it was measured. Because the height of the tide is
changing, the time of the reading is important. Estimate the place on the beach where the water level
would be without the waves, the still water level. There is no need to measure how far up the beach
the swash is going at the time of the measurement.
17. Continue On (Optional). The process can be continued into the water if teams want to. This is
optional and not necessary. In cold water there is a risk of hypothermia. In rough seas there is a risk
of getting hit by breaking waves. Do not take chances. Always keep your personal safety and that of your
team members in mind. A few extra points on a graph are not worth the risk of personal injury.
18. Final Reading. At the last measurement, record the TIME finished on the log sheet.
19. Photograph the Beach. Take three photographs of the beach. It helps to place the profile rods
down on the profile line part way up the beach, near the high-tide line. Stepping back from the rods,
take a picture looking up to the dune (or seawall) from a spot near the water line. Move up about
halfway on the profile and take two more pictures: one looking each way along the beach (parallel to
the water line). For these shots try and include the profile rods in the foreground. Frame the picture
to include the beach from dune
(seawall) to the water.
20. Pack Up. Gather up all the field
gear, including notes and any posts
back at the control points.
Using a home-made Emery rod to measure beach slope. Photo credit: Florida Sea Grant
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Longshore Current Measurement
Longshore currents affect shorelines by redistributing sand and sediment along their path. This
redistribution is also known as littoral drift. Longshore currents form because waves are continuous and, in
most cases, approach the shore at an angle. When a wave enters shallow water it is slowed by the rising
sandy or rocky bottom that is rising upward making a shallow edge. This friction eventually causes breakers
(the water you see toppling over at the water’s edge). As one wave meets the shore and breaks, another wave
is right behind it, preventing the broken wave from flowing backward. This causes a “build-up” of water at
the shoreline. This “build-up” of water is then forced to form a current that flows parallel to the shore close
to the water’s edge.
Equipment: Tape measure to measure off 10 meters, at least two oranges or pieces of driftwood; a
stopwatch or a wristwatch with a second hand.
Procedure:
1. Measure and mark off a 10-meter long line parallel to the ocean.
2. Determine which direction the longshore current is flowing by tossing an orange into the water just
beyond the breakers. Observe to see which direction the orange drifts.
3. Post one person each at the beginning and end of the 10-meter line and instruct them to look
straight ahead towards the water. Supply the person at the beginning of the line (the direction from
which the current is coming) with a stopwatch or a wristwatch with a second hand.
4. Go to the water’s edge slightly past the beginning of the line and toss an orange or piece of
driftwood into the breakers (just behind the white foam at the top of the wave). When the orange
passes the start line the person there should start timing.
5. The person at the other end should signal the timekeeper as soon as the orange passes his or her
post. The time is then recorded and the data sheet (page 79) is used to calculate the speed or the
velocity of the longshore current.
(Source: The Education Program at the New Jersey Marine Sciences Consortium)
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COASTAL FLORIDA ADOPT-A-WETLAND
PHYSICAL/CHEMICAL WATER MONITORING FORM
AAW Group Name County
Group ID Site ID
Investigators
Wetland Name Date Time
Site Location Description (e.g. Beach, Mangroves, Salt Marsh)
Rain in Last 24 Hours? Yes/No Amount of Rain Inches in Last hours/days □Heavy Rain □Steady Rain □Intermittent Rain
Present Conditions: □Heavy Rain □Steady Rain □Intermittent Rain □Partly Cloudy □Overcast □Clear/Sunny
Is Waterway Influenced by Tides? Yes/No If Yes, Tide was: □High □Outgoing □Low □Incoming Water Surface Conditions: □Calm □Ripples □Waves □White caps Impaired Habitat Indicators: □Foam □Bubbles □Oil □Scum □Dead Organisms □Vegetative Debris □Erosion □Dumping □Trash Present □Excessive Algae Water Color: □Clear □Muddy □Milky Gray □Green □Brown
□Tan □Other Odor: □Gas □Oi □Chemical □Other
Physical and Chemical Tests
Basic Tests: Sample 1 Sample 2 Average
Air Temperature _________ _________ (°C/°F) _________(°C/°F) Water Temperature _________ _________ (°C/°F) _________(°C/°F) Sampling Depth _________ _________ (cm) _________(cm) pH _________ _________ _________ Dissolved Oxygen _________ _________ (mg/L) _________(mg/L) Water Salinity _________ _________ (ppt) _________(ppt) Secchi Disk Depth _________ _________ (cm) _________(cm) Settleable Solids _________ _________ (ml/L) _________(ml/L)
AdditionalTests,ObservationsandComments:
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LONGSHORE CURRENT DATA SHEET GROUP ID: ______________________ SITE ID: _______________________________ TODAY'S DATE:___/___/___TIME OF DAY: _______TIDAL STAGE:____________ WEATHER CONDITIONS: ________________________________________________ WIND DIRECTION: __________ OBSERVATIONS: ___________________________ ________________________________________________________________________ Trial 1 Trial 2 Trial 3 Average
Distance (Length of Transect Line)
10 Meters 10 Meters 10 Meters
Distance of line in Feet Hint: 1 meter = 3.28 Feet
Time (in Seconds)
Speed of Current in Meters (speed = distance/time)
Speed of Current in Feet (speed = distance/time)
Direction of Current
NOTES:
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DATA SHEET FOR BEACH PROFILING Date: _____________ Start time: _______ am/pm Finish time: _______am/pm Location (GPS): N_________________ Tide state: __________________________ W_________________ Description of Location: _________________________________________________________ _____________________________________________________________________________ _____________________________________________________________________________ Position of back rod
Position of front rod
Difference in height (+/-)
Position of back rod
Position of front rod
Difference in height (+/-)
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WetlandWatcher
DeadorDyingMarsh/Swamp
Pollution
MarineDebris
DerelictTraps&Vessels
Microplastics
InvasiveSpecies
WildlifeViolations
HabitatEnhancementProjects
“WhoToCall”List
MakeYourOwnFieldEquipment
ChapterSixProblemsinYourAdoptedWetland?
Balloon debris in the wrack at Bill Baggs Cape Florida State Park. Photo credit: Maia McGuire
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Wetland Watcher
As a "wetland watcher," part of monitoring your adopted site is to be aware of natural and unnatural
changes to the wetland. When observing, take notes and look for browning, thinning, or die off of marsh
grasses or mangroves. Non-seasonal changes may indicate signs of stress. Other important problems to be
aware of and able to identify include pollution, such as oil or sewage, marine debris, invasive species, and
wildlife offenders.
Dead or Dying Vegetation
If your adopted wetland begins to show signs of stress, be sure to document them with photos and increase
your monitoring frequency. If the wetland continues to degrade, contact your local Sea Grant extension
agent. Wetland condition and appearance are important; over time and after many visits to your adopted
wetland site, you will become accustomed to its appearance and its condition in a healthy state. Over time, if
your data have any unusual changes or the visual appearance of your site changes, you will notice. There are
many factors that may affect your estuary such as weather, insects, disease, nutrients, pollution, predation,
and climate.
Salt marsh grasses tend to turn brown in the winter—this is a natural seasonal change. If the browning
continues through the spring and summer, there may be a reason to be concerned. Watch for die off of
marsh grass that leads to open muddy patches in the marsh. If the muddy patches continue to grow in size
there may be a problem that will need to be identified.
Mangroves also respond to stress, however changes related to stress are chemical and are generally not
visible. Temperatures below freezing for extended periods of time can kill mangroves. This is mainly
expected to be seen along the ecotone at which mangrove swamp transitions to salt marsh in northern
Florida. When exposed to freezing temperatures, mangrove leaves will turn black and fall from the tree. The
leaves of some mangrove species turn yellow and then are shed from the tree. This is a natural process and
should not raise concern.
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If your wetland shows signs of die-off be sure to contact your local Adopt-A-Wetland coordinator so that
further testing at the site can be done and the appropriate agencies notified.
Pollution
Pollution can be a main contributor to wetland die-off. Pollution is any substance introduced to the
environment that has harmful effects. Pollutants enter estuaries by different sources and are called point
source pollution and nonpoint source pollution. Point source pollution comes from a specific source that
can be identified such as a pipe, channel, or other obvious discharge point from a single location. Examples
of point source pollution are industrial discharge or wastewater from a treatment plant. Currently, the 1972
Clean Water Act regulates point source pollution. Nonpoint source pollution is much harder to regulate
because it comes from a variety of discharge areas rather than a single identifiable source. Examples of
nonpoint sources include storm water runoff from urban areas, marinas, agriculture, forestry, construction,
leaky septic tanks, vehicles, and lawns. Leaky septic tanks and pet waste that is not picked up pollute
waterways with harmful bacteria and excess nutrients that pose major health concerns. Sediment in runoff
from construction sites, agricultural activities, forestry operations, and dredging can carry harmful
environmental pollutants and increase turbidity. The sediments transported by the runoff can cover and kill
critical benthic habitats and also carry dangerous heavy metals and other pollutants with them. If you
identify a problem while monitoring, go to your contact list and choose an appropriate person/agency to
call.
Marine Debris
The National Oceanic and Atmospheric Administration defines marine debris as "any persistent solid
material that is manufactured or processed and directly or indirectly, intentionally, or unintentionally,
disposed of or abandoned into the marine environment or the Great Lakes." Marine debris poses a potential
threat to human health, endangers wildlife, and degrades habitat.
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There are many sources of marine debris, some from land and some from water. Land-based sources
include illegal dumping, litter, released balloons, and disposable items. Some sea-based sources include lost
commercial and recreational fishing gear, and shipping containers from cargo ships. Although most marine
debris is accidently introduced to the environment, some is intentionally dumped.
Plastic debris can last in the environment for hundreds of years. Plastics in the marine environment range
from monofilament fishing line to plastic bags, straws, and bottles to microplastics. Wildlife commonly
mistake plastics for food and ingest or become entangled. For example, sea turtles commonly mistake plastic
bags and balloons for jellyfish and eat them. The plastics cannot be digested, causing an intestinal blockage
(called impaction) which usually leads to starvation. Entanglement in lost or discarded fishing gear is
common. In sea turtles and marine mammals, entanglement often leads to lost appendages or death.
Coastal cleanups are an easy and effective way to protect wildlife and improve degraded habitats. Ask your
local Adopt-A-Wetland coordinator how to set up and conduct a coastal cleanup.
Derelict Traps
Derelict traps include lost or abandoned spiny lobster traps, stone crab traps, and blue crab traps.
Commercial and recreational traps become derelict when the owners can no longer locate them. Traps can
be moved by storms or have their floats cut off by boat propellers, making them difficult to relocate. These
lost or abandoned traps have been given the nickname "Ghost Traps" because they continue to trap
crustaceans, fish, and occasionally diamondback terrapins (Malaclemys terrapin) after they are no longer fished.
Once trapped in ghost traps the animals cannot escape, ultimately becoming bait themselves. Derelict traps,
discarded fishing gear, and marine debris pose numerous threats to marine wildlife. By removing these
items, our coastal wetlands are a much healthier and safer environment for marine wildlife.
It is a criminal offence to take or move a commercial or recreational trap (even if it is derelict) without a
permit from the Florida Fish and Wildlife Conservation Commission (FWC). Biannually, regional blue crab
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fisheries are closed by FWC for a short period to give governmental, private, and non-profit groups the
opportunity to remove derelict traps.8
Derelict Vessels
Florida's waterways are littered with abandon and derelict vessels that pose environmental and public safety
risks. The cost of removing the vessels can be very high, and the removal can be difficult, especially when
the vessels are abandoned on critical habitat. The vessel owner is responsible for all costs, but more often
than not it is virtually impossible to identify or locate the owner. In that case, the expense becomes a state
responsibility.
A main concern related to derelict vessels is public safety. Derelict vessels cause navigation issues and safety
concerns for boaters, especially at night when there are no lights on the derelict vessel. Derelict vessels left
adrift can cause extensive damage to seagrasses and other essential habitats. FWC's at risk vessel program is
working with boat owners whose vessels are showing signs of become derelict.9
Microplastics
Microplastics are defined as plastic or polymer particles that will fit through a 5mm mesh. Large-scale plastic
production became widespread in the 1950s. By the 1970s, scientists were starting to become concerned
about microplastics and their effects on the environment. Since then, these small floating plastic particles
have been discovered in water bodies around the world. The impacts these small plastics can have are
potentially huge in marine environments.
Today plastic is used in almost everything we use in one way or another, from kids’ toys to packing goods.
As a result, the amount of plastic entering the marine environment has greatly increased and plastic
pollution has become a global issue. Microplastics enter the marine environment from many sources.
8http://myfwc.com/fishing/saltwater/trap-debris/9http://myfwc.com/boating/waterway/derelict-vessels/
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Primary microplastics are those that originate as small particles, such as industrial scrubbers and micro beads
in personal care products. Secondary micro plastics are the result of larger items such as plastics bags,
bottles, and fishing lines breaking down as a result of ultraviolet light and chemical or microbial degradation.
Microplastics have the potential to pose threats to wildlife and human health. Toxic chemicals found in
water at very low concentrations have been found to adsorb to the surface of floating plastics. The
concentrations of these toxins on the plastic can be up to a million times higher than the levels found in
seawater. Additionally, toxic chemicals used in the manufacture of plastics (such as bisphenyl-A) can leach
from the plastic into organisms. The ingestion of microplastics by aquatic organisms can lead to health
concerns and possibly death. If microplastics move up the food chain and bioaccumulate, they could act as
vectors for toxic chemical transport.
To reduce microplastics in the environment, be aware of products that contain microbeads like toothpastes
and face washes. The simple act of not buying products that contain polyethylene plastic can potentially
reduce the amount of new microplastics in the marine environment. Additionally, recycling plastics,
reducing the use of single-use plastics (like water bottles, plastic cups, and straws), and reusing plastic
containers whenever possible can reduce the contribution to secondary microplastic pollution. To test your
adopted wetland for microplastics, sampling methods can be found on page (98).
Invasive Species
Invasive species are organisms that live outside of their native range and have a detrimental effect on natural
ecosystems, economies and/or human health. Invasive species often outcompete and displace native species
and can introduce new diseases and parasites. Common coastal wetland invasive species in Florida include
lionfish, green mussels, Brazilian pepper, and Australian pines.
The introduction of invasive species can be accidental; such as the discharge of ballast water containing
larval stages of invasive species from large ships near coastal waters, or accidental transport of invasive
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species from an improperly cleaned boat hull or bilge water from another region. However, the release of
ornamental fish from home aquaria is becoming a very common pathway. For information on reporting
invasive species, go to http://myfwc.com/contact/report/exotic-species/ or call 1-888-IVE-GOT1 (1-888-
483-4681). Also check out the Early Detection and Distribution Mapping System,10 an invasive species
tracker.
Lionfish:theFirstInvasiveMarineFishtobecomeEstablishedintheAtlantic
Indo-Pacific lionfish (Pterois volitans and P. miles) are the first reported non-native marine fish to become
established in the Atlantic Ocean. Their introduction was most likely the result of accidental or deliberate
release of aquarium pets. Currently in the United States, the lionfish is almost continuously distributed in
marine waters from the northern Gulf of Mexico to Cape Hatteras, North Carolina. In the wake of their
rapid and successful establishment in coastal waters of the southeast United States and greater Caribbean
region, there is concern that lionfish compete with and eat native fish that are ecologically and commercially
important. Several agencies and environmental groups have programs that allow and encourage the public to
help monitor and remove lionfish. You can report your catch to FWC.11 The Reef Environmental
10http://eddmaps.org/florida/ 11http://myfwc.com/reportlionfish
Invasive lionfish in the Atlantic. Photo credit: Kay Wells
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Education Foundation, or REEF, has been actively working in the Caribbean and Florida Keys for the past
several years to train divers to identify and safely catch and handle lionfish. In partnership with local
governments, REEF sponsors lionfish rodeos, which can result in the capture of over 1000 lionfish in a
single day.12
To learn more about Lionfish, their impacts to native species, and what you can do at the University of
Florida’s IFAS Extension website13 or visit the Lionfish Web Portal.14
Wildlife Violations15
When in the field, if you witness a wildlife violation, do not confront the violator. Stay safe, do not get
involved, go to a safe location and call 1-888-404-FWCC(3922) to report the violation to law enforcement.
Call 911 if there is concern for your safety. Examples of violations:
Illegal hunting
Taking illegal salt water species
Harassing protected species
Boating under the influence
Illegal dumping
Habitat Enhancement Projects
Habitat enhancement projects are designed to improve the health of your site. Examples of projects include
planting buffers, estuarine restoration, oyster recycling, and regular litter pick-ups. Some enhancement
projects may need to be permitted by the Department of Environmental Protection (DEP) and/or the U.S.
Army Corps of Engineers. Please check if permitting is needed before starting any enhancement projects.
12http://reef/org/lionfish13 http://edis.ifas.ufl.edu/sg132 14http://lionfish.gcfi.org 15http://myfwc.com/contact/wildlife-alert/
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WhoToCallList:
Florida Fish and Wildlife Conservation Commission Emergency Operation Center: Most or all
coastal emergencies including bird/fish kills, oil spills, pollution problems, whale/manatee/turtle
sightings/harassment/deaths: 888-404-FWCC (888-404-3922)
o Wildlife Law Violations: Call the Wildlife Alert hotline: Cellular phone users, call *FWC or
#FWC. Or send a text to [email protected]. : 888-404-FWCC (888-404-3922)
o Fish Kill Hotline: Call 800-636-0511, or submit a fish kill report to
http://myfwc.com/fishkill
o Report dead birds: http://legacy.myfwc.com/bird/default.asp
o Non-emergency wildlife issues: Contact your local regional FWC office.16
Nuisance Alligators: Call 866-FWC-GATOR (866-392-4286).
Orphaned, Injured or Nuisance Wildlife: Find local rehabilitator contacts at
http://myfwc.com/contact/nuisance-wildlife/
Oil, Fuel or Hazardous Material Spills in Florida waters: contact the Florida State Watch Office at
850-413-9900 (non-emergency) or 850-413-9911 or 800-320-0519 (emergency).
Physical/Chemical Pollution issues: Contact your local Department of Environmental Protection’s
district office.17
Red Tide Status Line: Toll-free inside Florida only 866-300-9399.Outside Florida 727-552-2448
Right Whale Sightings: Call 888-97-WHALE (888-979-4253).
Smalltooth Sawfish Sightings: 941-255-7403
Angler Tag Return Hotline: 800-367-4461.
Horseshoe Crab Nesting Activity: 866-252-9326
Florida Sea Grant: Main office 352-392-2801
16http://myfwc.com/contact/fwc-staff/regional-offices/17http://www.dep.state.fl.us/secretary/dist/
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MakeYourOwnFieldEquipment
Emery Rod Construction (From the Woods Hole Oceanographic Institution Sea Grant Program)
Emery Rod construction is quite simple. Necessary components include four PVC or wooden rods, each 1-
inch square or 1-inch diameter and 5 feet in length. Two rods are painted one side only with alternating
bright colors (such as red and white) tape can be used as well, with each block in increments of tenths of
feet, or in inches or centimeters, beginning at the top. The remaining two rods are connected to the painted
rods (with bolts and wingnuts) to form a parallelogram.
NOTE: To prevent the rods from sinking into the sand and thus giving inaccurate elevation readings, it is
recommended that ‘foot pads,’ made of small discs of wood or large bottle caps, be attached to the bottom
of the two vertical rods.
P
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Make an Aquascope to Explore Tide Pools
Visiting the rocky shore offers an exciting look at ocean plants and animals in the place they call home. Though
tide pool creatures survive harsh conditions, they’re easily hurt or disturbed by human visitors. Using a
homemade aquascope, you can watch tide pool life right where it is and leave the animals in their tide pool
homes.
Materials
Large “No. 10” can or large coffee can with both ends removed, waterproof plastic tape, heavy rubber bands,
clear plastic bag or food wrap, black paint (optional)
Directions
1. Paint the inside of the can with black paint (optional but helps viewing).
2. Cover the top and bottom rim of the can with plastic tape to cover the sharp edges.
3. Stretch the plastic bag or food wrap TIGHTLY over the bottom of the can.
4. Secure the plastic bag or wrap against the can with one or more heavy rubber bands.
5. Seal the edges of the plastic against the can with waterproof tape if available.
Leave ocean animals in their homes. Most will die if pried from the rocks, and all of them need the oxygen
from seawater to breathe. Always return animals exactly as you found them. Replace any rocks or shells that
you turn over—they are homes for many animals.
(Source: Monterey Bay Aquarium www.montereybayaquarium.org)
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How to Make a Viewscope
For a basic viewscope, you will need:
4"-diameter PVC pipe, black inside
1 handle, 3-4” long
4 screws and nuts (to attach handle)
Cut a 2-3 section of 4"- diameter PVC pipe. If you can't find pipe with a black interior, paint the inside black. If
the pipe is shiny black inside, use sandpaper to rough up the interior. Attach handle about 6" from one end.
Optional refinement: Programs that monitor in choppy waters may want to modify their viewscopes by
adding a Plexiglas "window" at one end. This prevents water from coming up inside the tube and interfering
with visibility. Materials needed are:
4.5" -diameter Plexiglas disk
PVC coupling
Silicone rubber sealant
Glue the Plexiglas disk to the bottom of the tube, using silicone rubber sealant. Place a piece of PVC coupling
over that end of the tube (like a collar) and seal with the silicone sealant. Drill two small (1/8") holes in the side
of the collar so that air won’t be trapped in the open end of the coupling when you put the viewscope into the
water.
(Jeff Schloss, Coordinator of New Hampshire Lakes Lay Monitoring Program http://dipin.kent.edu/makedisk.htm)
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Making a Quadrat
Quadrats are relatively simple and inexpensive to make. You will need the following materials and supplies for
one 1 meter x 1 meter square quadrat.
20 feet of ½ or ¾” PVC pipe (there are different grades of pipe—look for the cheapest kind)
4 ea. ½ or ¾” PVC elbows (90°)
PVC cement
Hacksaw or PVC cutters
Drill with small bit (1/4” or so…)
String
Tape measure
Instructions:
1. Cut the PVC pipe into four 39” pieces
2. Working in a well-ventilated area, use PVC glue to attach elbows to one piece of PVC. It is best to do
this on a flat surface, so you can make sure that the elbows are in the same plane. Use PVC glue to
attach the rest of the PVC together in a square.
3. Measure the center point of each side of the quadrat, and also mark the ¼ and ¾ points. Drill a hole
completely through the PVC at each of these points on each side. These holes will be used to attach
strings and also so that the quadrat will sink if used in water.
4. Tie the string through the holes from opposite side to opposite side so you end up with a grid that
divides the quadrat into 16 equal squares. You may need to burn the ends of the string to prevent it from
unraveling.
The inside dimensions of the finished product should be approximately 1 m (39.4”) on each side.
To make a smaller (1/2 m) quadrat, simply reduce the size of the PVC to the desired length.
Home-made quadrat. Photo credit: Maia McGuire
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HowtoMakeaSecchiDisk
A typical Secchi disk is 8” in diameter.
1. To make a Secchi disk, use a plastic lid from a small bucket
approximately 8” diameter. You can also cut down smaller lids
(using a Roto-zip; a jig-saw would also work) into 8” circles.
2. Use a marker to divide the lid into quarters, mask off two of the
quarters (diagonally opposite each other), then use black spray
paint to paint the remaining 2 quarters. Once the paint is
completely dry, carefully remove the masking tape.
3. Use a 4 oz fishing weight as the ballast for the Secchi disk and use a drill to make a hole in the center
of the disk and push the “eye” portion of the weight through the hole so that the “body” of the
weight is on the underside of the disk.
4. Attach a piece of thin rope to the eye of the weight. The knot in the rope prevents the weight from
separating from the plastic disk in the water.
To use the Secchi disk, lower it into the water until the
distinction between black and white sections disappears. Mark
the rope at that point (at the water’s surface). Retrieve the disk
and measure the distance from the mark to the top of the Secchi
disk. Lower the disk back into the water beyond the first mark,
then slowly pull it back up until you can just make out the
difference between the black and white sections. Make a new
mark on the rope and measure this second distance. The Secchi
depth is the average of the two numbers.
Home-made Secchi disk. Photo credit: Maia McGuire
US Environmental Protection Agency
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HowtoMakeaHester‐DendySampler
A Hester-Dendy sampler is a device which allows one to collect samples of algae, mollusks or other aquatic
organisms that attach themselves to underwater surfaces. You will need to leave this sampler in the same
location, undisturbed, for at least a few weeks.
Materials Needed:
Four or five pieces of scrap wood or plywood, roughly 4" square or round.
One eye bolt (about 10-12" long)
Two nuts and 2 washers for each plate of wood used
Lead weights (fishing weights will do nicely)
Cord to suspend sampler
What to Do:
1. Drill a hole in the center of each piece of wood.
2. Slip a piece of wood onto the eye bolt, holding it in place with a nut and washer on top and another
on the bottom of the wood.
3. Do the same with the other pieces of wood, making sure that there is a quarter to half inch between
each in order to give the organisms room to attach and grow.
4. Attach the lead weights to the bottom of the sampler. Use enough weights to keep the sampler still
while it is hanging in the water, or partially buried in a stream bottom.
5. Attach the cord to the eye of the bolt.
6. Tie the other end to a dock or pier where you know your sampler will be safe from tampering or
theft. If using the Hester-Dendy Sampler in a stream, tie it to a tree so it will not be swept
downstream by high water currents.
7. Come back in a few weeks and collect your samples.
8. Disassemble the sampler to facilitate viewing of the organisms which colonize it. Unscrew the nuts
on the eye bolt and gently swish the wood pieces in a pan of water to release the organisms which
have attached themselves to the surface.
(Source: http://www.esu7.org/~waterqweb/Macro_inverts/equipment.htm)
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Make Your Own Plankton Net
Plankton are very common in all bodies of water, but they are often very spread out and it would be hard to
look at them without making a "plankton concentrate". A net is the tool that is most often used to increase
the number of plankton within a volume of water. The resulting sample is then observed with a microscope
(but you can use a magnifying glass or just your eyes). A "professional" plankton net is comprised of a very
expensive nylon mesh with highly accurate hole sizes in it (mesh), but you do not need to be so complicated.
As long as a mesh can filter the critters out of the water, the net will accomplish its purpose. Here are
instructions showing how to construct a simple plankton net of your own out of common household items.
Materials needed:
A pair of nylon stockings (ask for a pair from a female member of your family - Do not just find
some - I guarantee trouble if you cut up your mom's good stockings!)
Wire coat hanger
Pair of pliers
Small (preferably plastic) bottle with a fairly small mouth size (an old pill bottle, small jar -
something with a lip around the mouth edge)
Scissors
Stapler or Duct tape
Rubber band (a medium wide one if you can find it)
A washer, a plastic ring, or long tie-tape
Strong string (kite string) or fine nylon twine.
Net Assembly: Cut one of the legs off of the nylons near the top. Unwind the coat hanger (be careful - you
might need some help with this), then create a ring about 15-25 cm (6-10 inches) in diameter (it can't be
much larger than the top of the cut stocking leg). Twist the ends of the hanger together with the pliers and
shape the wire into a circle as best as you are able. Put the top end of the leg through the wire ring and fold
it back over the outside of the wire ring. Staple or tape the nylon leg to attach it to the wire ring. Cut a small
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hole in the toe end of the nylons about the same size as the bottle mouth opening you have. Stretch the end
over the mouth of the bottle or jar. Wrap the rubber band tightly around the nylons to secure it to the
bottle.
You now need to make a pyramid of string in front of the net so the net will tow with the mouth of the net
facing forward and not collapse. To do this, you need to tie three evenly spaced strings, each about 60 cm (2
feet) long to the coat hanger ring at the top of the net. Tie the other end of these strings to a washer or
another small ring (you can use a tie-tape, but it will be harder to attach your towing rope to it) which will be
in front of the net when you tow it.
Plankton Collection: Attach a longer towing string or rope to the ring and tow it from a canoe or
motorized boat (go slowly and do not get it near the propeller), a dock, toss it (or drag it while wading) into
a pond or lake and pull it back a couple of times. The longer you tow, the more you will catch. To retrieve
the sample, remove the rubber band and dump your bottle into a tray (with dark color if you want to see
your plankton) or larger bottle to store the sample. You may see little dots swimming around, or possibly
just drifting. Plankton will not keep for long, so do not wait too long before looking at it (refrigeration will
help to keep it fresh, but not much longer than a day or two). You can also preserve the plankton by mixing
your water half-and-half with rubbing alcohol (ask someone responsible before using this - it is a dangerous
chemical). Unfortunately, most plankton are extremely small so you might not get much detail using just
your eyes (but many zooplankton are about 1 mm long so you can see them if you look carefully for them
swimming). A microscope is the most effective way to view your samples. If you do not have a microscope
available, try putting your sample into a large thick glass jar and tilt the jar around a little. The curvature of
the glass will cause some magnification of the things near the edge (check some of your jars at home to find
a good "magnifier" candidate).
(Source: http://www.biosci.ohiou.edu/faculty/currie/ocean/makeanet.htm)
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Sampling for Microplastics
Microplastics can be found in the sediment and in the water column. Here are some simple methods for
investigating this type of pollution.
A. In the sediment
Materials needed: Quadrat (1’ x 1’), container to hold sand (gallon zipper-seal bag, sealable bowl or tub),
small trowel or large spoon, paper plate(s), sieve (window-screen size, or 0.25 mm if using graded sand
sieves), tweezers, large cups (2-3), water
Procedure:
1. At the field site, randomly place the quadrat in the area of the wrack line.
2. Use the trowel/spoon to scrape about the top ¼ to ½ inch of sediment/wrack and scoop it into a
container or bag. Seal the container.
3. Indoors, pour the contents of the container onto paper plates and spread out the sediment to dry.
Leave at least overnight. If the sediment is already dry, you can skip this step.
4. Sift the sediment through the sieve. Capture the fine sand that comes through the sieve and save it
to return it to the field location.
5. Visually look through the sediment and debris left in the sieve (you can pour it back onto a clean
paper plate to help with this step.) Look for any obvious pieces of plastic and pick them out. Set
them aside in a small container.
6. Take the remaining sediment/debris and pour it into one or more large cups. Fill the cups about ¾
full with tap water. Stir well. If you see plastic pieces rise to the surface immediately, go ahead and
pick them out and add them to the ones previously found.
7. If there is plant material in the debris, it will also float (as will pieces of crab shell and small snail
shells that have air trapped in them). The longer the plant material soaks, the more likely it will be to
sink. If possible, leave the cups overnight and stir and check them again the next day before
discarding the contents.
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B. In the water
Materials needed: 1-liter bottles (any variety, but if purchasing, wide-mouth Nalgene bottles work well);
vacuum filter apparatus that can take 47-mm filters; 0.45 micron gridded filters; filter forceps; squirt bottle,
tap water; 1-liter separatory funnel and stand/clamp, dissecting microscope (20-30 or 20-40 X).
Procedure:
1. Triple-rinse 1-L bottles with water at your collection site. Be sure to discard your rinse water
downstream of where you/others are collecting samples. If using Nalgene bottles, you probably do not
need to be concerned about contamination by plastic from the threads, but if you are using other types
of plastic collection bottles, you should line the lids with foil.
2. Immerse the sample bottle sideways (holding it horizontally) into the water until it is just submerged.
Allow it to fill with water and cap it underwater.
3. You can let samples sit for weeks before processing (they do not need to be refrigerated, although they
should be kept in the dark to prevent algal growth in the bottles).
4. Run about 100 ml of tap water through a 0.45 micron filter (vacuum filter it). Use this to rinse the inside
of the side-arm flask (the one you've used to collect it in) and discard. Repeat 2 more times. (Essentially
you are triple-rinsing the flask with filtered water). Similarly triple rinse a squirt bottle with filtered tap
water. Collect the next 500 ml of filtered water and use it to stock the squirt bottle. You will use this
filtered water for rinsing the funnel, etc.
Sediment/wrack samples soaking (left); microplastics found in beach sand (right): Photo credits: Maia McGuire
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5. This part is optional, but recommended (it will make the samples much easier to filter). When ready to
process, triple rinse a 1-L separatory funnel. Pour the sample into the funnel (supported by a clamp on a
heavy-duty stand). Let sample stand for at least a few minutes. Drain off the sand/silt from the bottom
of the sample into a cup (this will be discarded).
6. With no filter inserted, rinse the inside of the filter apparatus with pre-filtered water. Use a petri dish or
other flat object as a cover for the filter apparatus (only remove when adding more sample). This will
help reduce environmental contamination of the sample (e.g. by lint in the air).
7. Insert the filter (gridded) into the apparatus. Add sample to fill the filter funnel. Put remaining sample
back on the clamp and allow to further settle (keep the separatory funnel or sample bottle stoppered).
Drain sediment from the separatory funnel as needed.
8. With the cover over the filter funnel, vacuum filter the sample. Add more sample until it has all been
run through the filter. Rinse the sides of the filter funnel with a small amount of filtered water once your
sample has been entirely filtered.
9. Release the vacuum pressure. Remove the filter and place into a clean petri dish. Cover with the petri
dish lid. Remember to label the sample (either on the petri dish lid, or with a small strip of paper placed
inside the petri dish, but not on the filter).
10. Let the filter dry at least overnight before viewing under a microscope (not required, but it's easier to
differentiate plastics from plankton once the plankton have dried out somewhat. It's also easier to scan
without the reflection from the wet filter).
11. If processing several samples collected in the same general location one right after the other, you do not
need to rinse the separatory funnel or filter funnel in between...but should do so before switching
sample locations.
12. Observe the filter papers under a microscope at 20X magnification. Scan the filters systematically,
moving row by row to prevent double-counting or missing plastics. Plastic will generally be milky/white
or colored (not clear). Sand grains are easily mistaken for plastics. Many of the fibers seen on the filters
will be extremely small.
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Plastics on filters (grid size of filters is 3 mm x 3 mm)
Photo credits: Maia McGuire
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Plankton on filters Photo credits: Maia McGuire
Copepod Ostracod Polychaete worm
Zoea larva (crustacean) Copepod with egg sacs Bivalve larva
Copepod (top); snail larva (bottom)
Sand grains
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Coulombe, D. 1992. The Sea Side Naturalist. Fireside Publishing, New York, NY, 246 pp.
Greene, T. F. 2004. Marine Science Marine Biology and Oceanography. Second edition. Amsco School Publications, Inc. New York, New York. 623 pp.
Hackney C.T., M. Adams, and W.H. Martin. Biodiversity of the Southeastern U.S. 1992. John Wiley and Sons, Inc. New York, New York, pp. 615-746.
Hickman, C.P., L.S. Roberts, F.M. Hickman. 1984. Integrated Principles of Zoology. Seventh edition. Times Mirror/Mosby College Publishing, St. Louis, MO.
Johnson, S., H.O. Hillestad., S.F. Shanholtzer., and G.F. Shanholtzer. 1974. An Ecological Survey of the Coastal Region of Georgia. National Parks Service Scientific Monograph Series, No. 3, 233 pp.
Long, S.P. and C.F. Mason. 1983. Salt Marsh Ecology. Blackie and Son Limited, Bishopbriggs, Glasgow. 153 pp.
Meadows P.S. and J.I. Campbell . 1978. An Introduction to Marine Science. Blackie and Son Limited, England, 176 pp.
Mitsch, W.J. and J.G. Gosselink. 1986. Wetlands. Van Nostrand Reinhold Co. New York, New York, 537 pp.
O’Beirn, F., P.B. Heffernan, and R.L. Walker. 1994. Recruitment of Crassostrea virginica: A Tool for Monitoring the Aquatic Health of the Sapelo Island National Estuarine Research Reserve. Marine Technical Report. No. 94-2.
Odum, W.E., C.C. Mclvor, and T.J. Smith, III. 1982. The ecology of the mangroves of south Florida: a community profile. U.S. Fish and Wildlife Service, Office of Biological Services, Washington, D.C. FWS/OBS-81/24. 144 pp. Reprinted September 1985.
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Bibliography
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Appendices
MacroinvertebrateIdentificationKey
PlantIdentificationKey
FishIdentificationKey
CommonMollusksofFlorida
OtherCommonMarineInvertebratesofFlorida
CommonFishesofFlorida
InvasiveAquaticSpeciesinFlorida
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Key to Macroinvertebrates Found in Coastal Florida
OR OR
A. Soft body contained in a thick, hard shell (Phylum Mollusca)
B. Body not soft/not in a thick, hard shell (Phylum Arthropoda or Porifera) See pages 112 - 115
C. Soft body but not inside hard shell (Phyla Annelida/Echinodermata/ Cnidaria/Ctenophora/Chordata) See pages 116 - 119
Body inside a single shell (Class Gastropoda) See page 109
Body inside 2 equal shells (Class Bivalvia) See page 110-111
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Animals with segmented legs and obvious joints (Phylum Arthropoda)
Body is not soft or not in thick. hard shell
Odd-shaped body with small holes (pores). Skin is rough (Phylum Porifera) See page 115
Shell-like, oval shaped, with plates. Attached to debris, pilings or docks (Class Cirripedia, barnacles) See page 113
One pair of antennae, 3 pairs of legs. Class Insecta. See page 114
Class Pycnogonida (Sea spiders) See page 114
Class Merostomata (Horseshoe crabs) See page 114
2 pairs of antennae, more than 3 pairs of legs (Subphylum Crustacea) See page 112-113
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Flattened laterally (sideways) so it looks like the legs are located on one side of the body Order Amphipoda) See page 113
Body is not soft or not in thick. hard shell (Subphylum Crustacea)
Head is jointed, eyes and antennae movable. Order Stomatopoda (e.g. Mantis shrimp)
7 pairs of legs, body flattened (Order Isopoda)
5 pairs of legs; head and thoracic segments fused together forming thin shell; pinchers present in some (e.g. crabs, shrimp) See pages 112 - 113
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Soft body, elongated thin shape. Worm-like (Phylum Annelida) See page 116
Soft body not in a. hard shell
Soft-bodied, stout shape. Often attached to shell/dock/pilings. Tentacles with stinging cells on one end (e.g. Sea anemone, jellyfish). (Phylum Cnidaria or Ctenophora). See pages 118 - 119
Odd shape, skin rough & fleshy. Most have 2 openings. No stinging cells (e.g. sea squirts, sea pork). (Phylum Chordata, subphylum Urochordata) See page 119
Odd-shaped body, skin rough. Some covered in spines. Small opening (mouth) on bottom side. (e.g. sea cucumber, sea star) (Phylum Echinodermata) See page 117
Segmented flat worms with rounded anterior and posterior ends (suckers). (Class Hirudinea)
Segmented worms with small appendages & antennae (Class Polychaeta) See page 116
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Knobbed Whelk Busycon carica
Channeled Whelk Busycon canaliculatum
Auger Terebra dislocata
Phylum Mollusca Class Gastropoda
Olive snail Oliva sayana
Mud snail Ilyanassa obsoleta
Marsh periwinkle Littorina irrorata
Coffee bean snail Melampus bidentatus
Atlantic Oyster Drill Urosalpinx cinerea
Lightning Whelk Busycon sinistrum
Whelk egg string
Florida Horse Conch Pleuroploca gigantean
Rock shell Stramonita haemastoma
True tulip Fasciolaria tulipa
Moon snail Neverita duplicata
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Phylum Mollusca Class Bivalvia
`
Razor Clam/ Jacknife Clam Ensis directus
Northern Quahog Mercenaria mercenaria
Eastern Oyster Crassostrea virginica
Disk Clam Dosinia discus
Carolina Marsh Clam Polymesoda caroliniana
Alternate Tellin Tellina alternata
Angel Wing Cyrtopleura costata
Stout Tagelus Tagelus plebeius
Pen Shell Atrina rigida
Calico Scallop Argopecten gibbus
Southern Surf Clam Spisula ravaneli
Atlantic Bay Scallop Argopecten irradians
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Class Bivalvia Continued
a
Ponderous Ark Noetia ponderosa
Blood Ark Anadara ovalis
Giant Atlantic Cockle Dinocardium robustum
Turkey wing Arca zebra
Incongrous Ark Scapharca brasiliana
Eared Ark Anadara notabilis
American Horsemussel Modiolus americanus Atlantic Ribbed Mussels
Geukensia demissa
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Female Phylum Arthropoda Male Subphylum Crustacea
Thoracic Sternites Thoracic Sternites
Blue crab Callinectes sapidus
Spider crab Libinia dubia
Green Porcelain Crab Petrolisthes armatus
Calico Crab Hepatus epheliticus
Striped Hermit Crab Clibanarius vittatus
Stone Crab Menippe mercenaria Speckled swimcrab
Arenaeus cribarius
Wharf crab Sesarma cinereum
Mud crab Panopeus herbstiiMole crab
Emerita talpoida
Lady Crab Ovalipes ocellatus
Ghost Crab Ocypode cordimana
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Phylum Arthropoda Subphylum Crustacea
Fragile Barnacle Chthamalus fragilis
Brown shrimp Farfantepenaeus aztecus
White Shrimp Litopenaeus setiferus
Pink Shrimp Penaeus duorarum
Amphipod
Skeleton Shrimp Caprella penantis
Goose Barnacles Lepas anatifera
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Phylum Arthropoda
Class Merostomata Class Pycnogonida
Class Insecta
Dragonfly
Mayfly Nymph
Midge Fly Larva
Pycnogonid Sea Spider
Horseshoe Crab Limulus polyphemus
Damselfly
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Phylum Porifera
Finger Sponge Haliclona oculata
Redbeard Sponge Microciona prolifera
Vase Sponges Scypha sp.
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Phylum Annelida Class Polychaeta
Polychaete Worm
Polychaete Worm Head (close up)
Polychaetae worm tube
Class Hirudinea
Leech
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Phylum Echinodermata
Class Ophiuroidea Class Asteroidea
Smooth Brittle Star Common Sea Star Ophioderma brevispinum Asterias forbesi
Class Echinoidea
Sand Dollar or Keyhole Urchin Sea Urchin Mellita quinquiesperforata Lytechinus variegatus
Class Holothuroidea
Sea Cucumber Sclerodactyla briareus
Urchin test
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Phylum Cnidaria
Class Anthozoa
Class Scyphozoa
Sea Pansy Renilla reniformis
Brown Anemone Aiptasia pallida
Sea Whip
Leptogorgia virgulata
Cannonball Jellyfish Stomolophus meleagris
Sea Nettle Chrysaora quinquecirrha
Moon Jellyfish Aurelia aurita
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Phylum Ctenophora
Class Tentaculata
Comb Jellies
Phylum Chordata
Subphylum Urochordata
Rough Sea Squirts Sea Pork Styela plicata Aplidium constellatum
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Wetland Vegetation by Zones
Saltmarsh: Low Marsh Zone High Marsh Zone Marsh Border Upper Marsh Border & Transitional Zone
Mangrove Swamp
Beach: Upper Beach Zone Primary Dunes Dune Meadows
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Low Marsh Zone Vegetation
Smooth Cordgrass Spartina altemiflora
Smooth Cordgrass: broad leaf blade, plant size varies.
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High Marsh Zone Vegetation
Glasswort Salicornia virginica, S. bigelovii, S. europaea
Saltwort Batis maritima
Salt Grass Distichlis spicata
Glasswort or Pickle weed: succulent plant with tiny bract-like leaves.
Salt Grass: leaf blades in one plane, summer-fall (beach meadows).
Saltwort: succulent leaf, prostrate woody stem.
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Marsh Border
Marsh Aster Aster tenuifolius
Sea Oxeye Borrichia fruitescens
Needle Rush Juncus roemerianus
Orach Atriplex patula
Sea Lavender Limonium carolinianum
Marsh Aster: small sparsely arranged lavender or white aster flowers with yellow centers, fall. Needle Rush: long tubular leaves with sharp points, painful to walkers.
Sea Oxeye: succulent leaf, yellow aster flower, spiny burr, summer.
Sea Lavender: small sparsely arranged purple flowers, basal leaves, fall.
Orach: similar to orach on beaches (A. arenaria) but smaller leaves.
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Upper Marsh Border and Transition Zone
Red Cedar Juniperus virginiana
Marsh Elder Iva frutescens
Cabbage Palm Sabal palmetto
Saltcedar or Tamarisk Tamarix gallica
Saltbush or Groundsel-Tree Baccharis halimifolia
Red Cedar: short needles, blue berry-like cones, juniper tree.
Cabbage Palm: similar to saw palmetto of the forest but pinnately.
Marsh Elder: serrated leaves, tiny flowers or seeds at end of stems, leaves not as fleshy as beach elder.
Groundsel-tree or Cotton Bush: irregularly-shaped leaves, cotton-like seed tufts in the fall, shrub.
Saltcedar or Tamarisk: small tree or shrub, similar to red cedar but paler green and more delicate, tiny
pink flowers at tips of stems, summer. Non-native.
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Upper Beach Zone Vegetation
Orach Atriplex arenaria
Sea Rocket Cakile edentula
Beach Croton Croton punctatus
Orach: succulent gray-green leaf, red stem, summer.
Beach Croton: dusky gray-green leaves and stem, round fruit, spring.
Sea Rocket: succulent plant, two-section fruit, dies in summer, spring.
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123
Primary Dunes
Sea Oats Uniola paniculata
Bitter Panic Grass Panicum amarum
Sandspur Centrus tribuloides
Beach Elder Iva imbricata
Sea Oats: seed head on tall stalk, curly leaf blade, summer-fall.
Sandspur: prostrate, sharp painful burr, fall.
Beach Elder: succulent leaf, woody stem, summer.
Bitter Panic Grass: broad, alternate leaf blades on the stalk, summer.
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Sea-Purslane Sesuvium portulacastrum
Primary Dunes Continued
Fiddle-Leaf Morning Glory Ipomoea stolonifera
Railroad Vine Ipomoea pes-caprea
Russian Thistle Salsola kali
Fiddle-Leaf Morning Glory: succulent leaf, large with flower, vine, summer-fall.
Railroad Vine: large purple flower, vine, fall.
Russian Thistle: succulent leaf with spine, small prickly flowers, summer. Non-native.
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Dune Meadows
Camphorweed Heterotheca subaxillaris
Butterfly Pea Centrosema virginianum Clitoria mariana
Grass-Leaf GoldenAster Heterotheca graminifolia
Wild Bean Strophostyles umbrellata
Dune Primrose Oenothera humifusa
Camphorweed: yellow aster flower, fall.
Wild Bean: small red pea flower, slender pod, vine, summer-fall.
Butterfly Pea: large purple pea flower, vine, spring-fall.
Grass-Leaf Golden Aster: yellow aster flower, grass-like leaf, summer.
Dune Primrose: prostrate, pink and yellow flower, spring and fall.
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Mangrove Swamp
Red mangrove: Lower side of leaf green; propagules cigar-like
Black mangrove: Lower side of leaf greyish; often salt crystals on upper surface of leaves; propagules
lima bean-like
White mangrove: leaf tips blunt with a slight notch; two visible pores at base of leaves
Buttonwood: leaves less succulent than true mangroves; button-like fruit that breaks apart into seeds
Red mangrove Rhizophora mangle
Image credit: Johannes Zorn
(1739-1799)
Black mangrove Avicennia germinans Image credit: Klaus
Schonitzer-Scan
White mangrove Laguncularia racemose Photo credit: WD Brush
Buttonwood Conocarpus erectus
Photo credit: WD Brush
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Florida Fish Identification Key
ATLANTIC THREAD HERRING ATLANTIC MENHADEN Opisthonema oglinum Brevoortia tyrannus
ATLANTIC NEEDLEFISH AMERICAN HARVESTFISH Strongylura marina Peprilus paru
MOSQUITO FISH LADYFISH Gambusia affinis Elops saurus
STRIPED ANCHOVY BAY ANCHOVY Anchoa hepsetus Anchoa mitchilli
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PIGFISH GREY SNAPPER Orthopristis chrysoptera Lutjanus griseus
ATLANTIC BUMPER INSHORE LIZARDFISH Chloroscombrus chrysurus Synodus foetens
ATLANTIC CUTLASSFISH SILVER JENNY Trichiurus lepturus Eucinostomus gula
ATLANTIC SPANISH MACKEREL ATLANTIC BUTTERFISH Scomberomorus maculates Peprilus triacanthus
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TARPON CREVALLE JACK Megalops atlanticus Caranx hippos
SILVERSIDE POMPANO Menidia menidia Trachinotus carolinus
SPOT WHITING/KINGFISH Leiostomus xanthurus Menticirrhus americanus
SHEEPSHEAD LOOKDOWN Archosargus probatocephalus Selene vomer
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BLACK DRUM RED DRUM Pogonias cromis Sciaenops ocellata
STAR DRUM SILVER PERCH Stellifer lanceolatus Bairdiella chrysoura
BLACK SEA BASS SAND PERCH Centropristis striata Diplectrum formosum
ROCK SEA BASS OYSTER TOADFISH Centropristis philadelphica Opsanus tau
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SMOOTH PUFFER NORTHERN PUFFER Lagocephalus laevigatus Sphoeroides maculatus
STRIPED BURRFISH NORTHERN SEAROBIN Chilomycterus schoepfi Prionotus carolinus
STRIPED SEAROBIN BIGHEAD SEAROBIN Prionotus evolans Prionotus tribulus
GAFFTOPSAIL SEA CATFISH HARDHEAD SEA CATFISH Bagre marinus Ariopsis felis
PALESPOTTED EEL SHRIMP EEL Ophichthus ocellatus Ophichthus cruentifer
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STRIPED MULLET ATLANTIC SPADEFISH Mugil cephalus Chaetodipterus faber
SILVER SEATROUT SPOTTED SEATROUT Cynoscion nothus Cynoscion nebulosus
ATLANTIC CROAKER WEAKFISH Micropogonias undulates Cynoscion regalis
REMORA SPOTTAIL PINFISH Remora remora Diplodus holbrooki
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LINED SEAHORSE OPOSSUM PIPEFISH Hippocampus erectus Oostethus brachyurus
WINDOWPANE OCELLATED FLOUNDER Scophthalmus aquosus Ancylopsetta flounder
SUMMER FLOUNDER BLACKCHEEK TONGUEFISH Paralichthys dentatus Symphurus plagusia
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SOUTHERN FLOUNDER HOGCHOKER Paralichthys squamilentus Trinectes maculatus
COMMON SNOOK Centropomus undecimalis
CLEARNOSE SKATE SOUTHERN STINGRAY Raja eglanteria Dasyatis Americana
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ATLANTIC STINGRAY SMOOTH BUTTERFLY RAY Dasyatis sabina Gymnura micrura
SANDBAR SHARK Carcharhunus plumbeus
LEMON SHARK BONNETHEAD SHARK Negaprion brevirostris Sphyrna tiburo
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Common Mollusks of Florida Phylum: MOLLUSCA Class: GASTROPODA
Family Acmaeidae
Scientific Name Lottia antillarum
Common Name Keyhole Limpet
Calyptraeidae
Crepidula aculeata (Gmelin, 1791) Crepidula fornicata (Linnaeus, 1758)
Spiny Slipper Atlantic Slipper
Crepidula plana (Say, 1822) White Slipper
Columbellidae
Anachis avara (Say, 1822)
Greedy Dove Snail
Columbella rusticoides Common Dove Snail
Dentaliidae
Dentalium laqueatum (Verrill, 1885)
Panelled/Reticulate Tusk
Epitoniidae
Epitonium angulatum (Say, 1830) Epitonium humphreysii (Kiener, 1838) Epitonium rupicola (Kurtz, 1860)
Angulate Wentletrap Humphrey’s Wentletrap Brown-Band Wentletrap
Fasciolariidae
Fasciolaria hunteria (Perry, 1811) Fasciolaria tulipa (Linnaeus, 1758) Pleuroploca gigantea (Kiener, 1840)
Banded Tulip True Tulip Florida Horse Conch
Favorinidae
Cratena pilata
Striped Nudibranch
Littorinidae
Littorina irrorata (Say, 1822)
Marsh Periwinkle
Melampodidae
Melampus bidentatus (Say, 1822)
Common Marsh Snail
Melongenidae
Busycotypus canaliculatus(Linnaeus, 1758) Busycon carica (Gmelin, 1791) Busycon carica kieneri (Philippi, 1848) Busycon sinistrum (Hollister, 1958) Busycotypus spiratus (Lamarck, 1816)
Channeled Whelk Knobbed Whelk Kiener’s Whelk Lightning Whelk Pear Whelk
Muricidae
Eupleura caudata (Say, 1822) Muricanthus fulvescens (Sowerby, 1834) Phyllonotus pomum (Gmelin, 1791)
Rough Oyster Drill Giant Eastern MurexApple Murex
Muricidae
Thais haemastoma canaliculata (Gray, 1839) Thais haemastoma floridana (Conrad, 1837) Urosalpinxcinerea (Say, 1822)
Southern Oyster Drill Florida Rock Shell Atlantic Oyster Drill
Nassariidae Naticidae
Ilyanassa obsoleta (Say, 1822) Nassarius trivittatus (Say, 1822) Neverita duplicata (Say, 1822) Sinum perspectivum (Say, 1831)
Eastern Mudsnail New England Nassa Moon Snail/Shark Eye White Baby’s Ear
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Olividae Oliva sayana (Ravenel, 1834) Lettered Olive
Pyramidellidae
Olivella mutica (Link, 1807) Boonea impressa (Say, 1822)
Variable Dwarf Olive Impressed Odostome
Terebridae
Terebra dislocata (Say, 1822)
Common Eastern Augur
Class: BIVALVIA
Family Anomiidae
Scientific Name Anomia simplex (d’Orbigny, 1842)
Common Name Common Atlantic Jingle
Arcidae
Anadara lienosa floridana (Conrad, 1869) Anadara ovalis (Brugière, 1789) Anadara transversa (Say, 1822) Arca zebra (Swainson, 1833) Barbatia candida (Helbling, 1779) Barbatia domingensis (Lamarck, 1819) Noetia ponderosa (Say, 1822)
Cut-Ribbed Ark Blood Ark Transverse Ark Turkey Wing White-Bearded Ark White Miniature Ark Ponderous Ark
Cardiidae
Dinocardium robustum (Lightfoot, 1786) Laevicardium laevigatum (Linnaeus, 1758) Laevicardium mortoni (Conrad, 1830) Laevicardium pictum (Ravenel, 1861) Trachycardium egmontianum (Shuttleworth, 1856)
Giant Atlantic Cockle Common Egg Cockle Morton’s Egg Cockle Ravenel’s/Painted Egg Cockle Florida Prickly Cockle
Chamidae
Arcinella cornuta (Conrad, 1866)
Florida Spiny Jewelbox
Corbiculidae
Polymesoda caroliniana (Bosc, 1801)
Carolina Marshclam
Donacidae
Donax variabilis (Say, 1822)
Florida Coquina
Glycymerididae
Glycymeris americana (DeFrance, 1829)
Giant American Bittersweet
Lucinidae
Divaricella quadrisulcata (d’Orbigny, 1842) Linga pensylvanica (Linnaeus, 1758)
Cross-Hatched Lucin Pennsylvania Lucine
Mactridae
Raeta plicatella (Lamarck, 1818) Mactra fragilis (Gmelin, 1791) Mulinia lateralis (Say, 1822) Spisula raveneli (Conrad, 1831) Rangia cuneata (Sowerby, 1831) Spisula solidissima
Channeled Duckclam Fragile Surfclam Dwarf Surf Clam Southern Surf Clam Common Rangia Atlantic Surf Clam
Myidae
Sphenia antillensis (Dall & Simpson, 1901)
Antillean Sphenia
Mytilidae
Amygdalum papyrium (Conrad, 1846) Brachidontes exustus (Linnaeus, 158) Geukensia demissa (Dillwyn, 1817)
Paper Mussel Scorched Mussel Atlantic Ribbed Mussel
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Mytilidae Ischadium recurvum (Rafinisque, 1820) Lioberus castaneus (Say, 1822) Modiolus americanus (Leach, 1815)
Hooked Mussel Chestnut Mussel Tulip Mussel
Ostreidae
Crassostrea virginica (Gmelin, 1791) Ostrea equestris (Say, 1834)
American Eastern Oyster Crested Oyster
Pandoridae
Pandora trilineata (Say, 1822)
Say’s/Threeline Pandora
Pectinidae
Aequipecten muscosus (Wood, 1828) Argopecten gibbus (Linnaeus, 1758) Nodipecten nodosus (Linnaeus, 1758) Pecten ziczac (Linnaeus, 1758) Chlamys sentis (Reeve, 1853)
Rough Scallop Atlantic Calico Scallop Lion’s Paw Zigzag Scallop Sentis/Scaly Scallop
Pholadidae
Barnea truncata (Say, 1822) Cyrtopleura costata (Linnaeus, 1758) Martesia cuneiformis (Say, 1822)
Fallen Angel Wing Angel Wing Wedge Piddock
Pinnidae
Atrina rigida (Lightfoot, 1786) Atrina seminuda (Lamarck, 1819) Atrina serrata (Sowerby, 1825)
Rigid Penshell Half-naked Penshell Saw-toothed Penshell
Pteriidae
Pteria colymbus (Röding, 1798)
Atlantic Wing Oyster
Semelidae
Abra aequalis (Say, 1822) Cumingia tellinoides (Conrad, 1837) Semele proficua (Pulteney, 1799) Semele purpurascens (Gmelin, 1791)
Atlantic Abra Common Cumingia White Atlantic Semele Purplish Semele
Solecurtidae
Tagelus divisus (Spengler, 1794) Tagelus plebeius (Lightfoot, 1786)
Purplish Tagelus Stout Tagelus
Solenidae
Ensis directus (Conrad, 1843) Solen viridis (Say, 1821)
Atlantic Jackknife Clam Green Jackknife Clam
Spondylidae
Spondylus americanus (Hermann, 1781)
Atlantic Thorny Oyster
Tellinidae
Macoma balthica (Linnaeus, 1758) Macoma constricta (Bruguière, 1792) Tellina alternata (Say, 1822) Tellina listeri (Röding, 1798)
Baltic Macoma Constricted Macoma Alternate Tellin Speckled Tellin
Veneridae Chione intapurpurea (Conrad, 1849) Lady-in-Waiting Venus
Chione latilirata (Conrad, 1841) Imperial Venus Dosinia discus (Reeve, 1850) Disk Clam Macrocallista maculata (Linnaeus, 1758) Calico Clam Macrocallista nimbosa (Lightfoot, 1786) Sunray Venus Mercenaria campèchiensis (Gmelin, 1791) Southern Quahog Mercenaria mercenaria (Linnaeus, 1758) Northern Quahog Mercenaria mercenaria (ecolocgical form notata ) Northern Quahog
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Other Common Marine Invertebrate Organisms of Florida Phylum: PORIFERA Class: DEMOSPONGIAE (Sponges)
Family Microcionidae
Scientific Name Microciona prolifera
Common Name Redbeard Sponge
Desmacidonidae Clionidae
Haliclona oculata Cliona Sp.
Finger Sponge Boring Sponge
Homocoelidae
Scypha Sp.
Basket Sponge Phylum: CNIDARIA Class: ANTHOZOA (Anemones)
Gorgoniidae Leptogorgia virgulata Sea Whip
Renillidae Actinidae
Renilla reniformis Actinoporus elegans
Sea Pansy Brown-Striped Anemone
Class: SCYPHOZOA (Jellyfish)
Stomolophidae Stomolophus meleagris Cannonball Jellyfish
Ulmaridae Pelagidae
Aurelia aurita Chrysaora quinquecirrha
Moon Jellyfish Sea Nettle
Phylum: CTENOPHORA Class: TENTACULATA (Comb Jellies) Bolinopsidae Mnemiopsis mccradyi Comb Jelly
Phylum: ECHINODERMATA Class HOLOTHUROIDEA (Sea Cucumbers) Sclerodactylidae Sclerodactyla briareus Sea Cucumber Class: ASTEROIDEA (Sea Stars) Asteriidae Asterias forbesi Sea Star
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Phylum: ANNELIDA Class: POLYCHAETA (Worms)
Nereidae Nereis succinea Southern Clam Worm
Onuphidae Sabellidae
Diopatra cuprea Fabricia sabella
Plumed Worm Fan Worm
Class: ECHINOIDEA (Sea Urchins, Sand Dollars)
Mellitidae Mellita quinquiesperforata Key Hole Urchin
Toxopneustidae
Lytechinus variegates
Sea Urchin Class: OPHUROIDEA (Brittle Stars) Ophiodermatidae Ophioderma brevispinum Smooth Brittle Star Phylum: ARTHROPODA Class: PYCNOGONITA (Sea Spiders) Tanystylidae Tanystylum orbiculare White Sea Spider Class: CIRRIPEDIA (Barnacles)
Cthamalidae Chthamalus fragilis Fragile Barnacle
Balanidae Lepadidae
Balanus eburneus Lepas anatifera
Ivory Barnacle Goose Neck Barnacle
Class: MEROSTOMATA (Horseshoe crabs) Limulidae Limulus polyphemus Atlantic Horseshoe Crab
Class: MALACOSTRACA (Crabs, Shrimps)
Portunidae Callinectes sapidus Blue Crab
Xanthidae
Calappidae
Panopeus obesus Menippe mercenaria Rhithropanopeus harrisii Hepatus epheliticus
Mud Crab Stone Crab
White-Fingered Mud Crab Dolly Varden/Calico Crab
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Leucosiidae Majidae
Persephona punctata Libinia emarginata
Purse Crab Common Spider Crab
Portunidae
Carcinus maenas Ovalipes ocellatus Arenaeuscribarius
Green/Porcelain Crab Lady Crab Speckled Crab
Grapsidae
Sesarma reticulatum Sesarma cinereum
Marsh Crab Wharf Crab
Ocypodidae
Uca pugnax Uca minax
Mud Fiddler Crab Brackish Fiddler Crab
Ocypodidae
Uca pugilator
Sand Fiddler Crab
Penaeidae
Penaeus aztecus Penaeus duorarum Penaeus setiferus
Brown Shrimp Pink Shrimp White Shrimp
Hippolytidae
Hippolyte sp.
Grass Shrimp
Squillidae
Squilla empusa
Mantis Shrimp
Diogenidae
Clibanarius vittatus
Striped Hermit Crab
Hippidae
Emerita talpoida
Mole Crab
Gammaridae
Gammarus palustris
Scud Amphipod
Haustoriidae
Haustorius Canadensis
Digger Amphipods
Caprellidae
Caprella equilibra
Skeleton Shrimp Phylum: CHORDATA Class: ASCIDIACEA (Tunicates, Sea Squirts) Styelidae Styela plicata Rough (pleated) Sea Squirts Molgulidae Molgula manhattensis Sea grapes Polyclinidae Aplidium constellatum Sea Pork
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Common Fishes of Florida Phylum: CHORDATA Class: OSTEICHTHYES (Bony Fishes) Family Acipenseridae
Scientific Name Acipenser brevirostrum
Common Name Atlantic Sturgeon
Lepisosteiformes Lepisosteus osseus Longnose Gar
Elopidae
Elops saurus Megalops atlanticus
Ladyfish Tarpon
Anguillidae
Auguilla rostrata
American Eel
Ophichthidae Ophichthus gomesi Ophichthus ocellatus
Shrimp eel Palespotted eel
Clupeidae Opisthonema oglinum Brevoortia tyrannus
Atlantic Thread Herring Atlantic Menhaden
Engraulidae
Anchoa hepsetus Anchoa mitchilli
Striped Anchovy Bay Anchovy
Ariidae
Arius felis Bagre marinus
Sea Catfish Gafftopsail Catfish
Synodontidae Synodus foetens Inshore Lizardfish
Gadidae Urophycis regia Urophycis foridana
Spotted Hake Southern Hake
Batrachoididae Opsanus tau Oyster Toadfish
Gobiesocidae Gobiesox strumosus Skilletfish
Belonidae Strongylura marina Atlantic Needlefish
Cyprinodontidae Cyprinodon variegates Fundulus majalis
Sheepshead minnow Striped Killifish
Poeciliidae Gambusia affinis Poecilia latipinna
Mosquitofish Sailfin Molly
Atherinidae Menidia menidia Atlantic silverside
Syngnathidae Hippocampus erectus Oostethus brachyurus
Lined Seahorse Northern pipefish
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Triglidae Prionotus carolinus Prionotus evolans Prionotus tribulus
Northern Sea Robin Striped Sea Robin Big Head Sea Robin
Bothidae Ancylopsetta quadrocellata Scophthalmus aquosus Paralichthys dentatus Paralichthys squamilentus
Ocellated Flounder Windowpane Summer Flounder Southern Flounder
Soleidae Trinectes maculates Symphurus plagiusa
Hogchoker Blackcheek Tonguefish
Balistidae Monacanthus hispidus Planeheaded Filefish
Ostraciidae Lactophrys quadricornis Scrawled Cowfish
Tetraodontidae Lagocephalus lagocephalus Sphoeroides maculates Chilomycterus schoepfi
Smooth Puffer Northern Puffer Striped burrfish
Centropomidae Centropomus undecimalis
Common snook
Percichthyidae Morone saxatilis Striped Bass
Serranidae Centropristis Philadelphia Centropristis striata Diplectrum formosum Mycteroperca phenax Mycteroperca microlepis
Rock Sea Bass Black Sea Bass Sand Perch Scamp Gag
Pomatomidae Pomatomus saltatrix Bluefish
Echeneidae Remora remora Sharksuckers
Carangidae Caranx hippos Chloroscombrus chrysurus Selene vomer Trachinotus carolinus
Crevalle Jack Atlantic Bumper Lookdown Florida Pompano
Lutjanidae Lutjanus griseus Gray Snapper
Gerreidae Eucinostomus gula Silver Jenny
Haemulidae Orthopristis chrysoptera Pigfish
Sparidae Archosargus probatocelphalus Diplodus holbroooki
Sheepshead Pinfish
Sciaenidae Bairdiella chrysoura Cynoscion nebulosus
Silver Perch Spotted Seatrout
Sciaenops ocellatus Leiostomus xanthurus
Red Drum Spot
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Larimus fasciatus Pogonias cromis Menticirrhus americanus Menticirrhus littoralis Micropogonias undulates Stellifer lanceolatus
Banded Drum Black Drum Kingfish/Whiting Gulf Kingfish Atlantic Croaker Star Drum
Ephippidae Chaetodipterus faber Atlantic Spadefish
Chaetodontidae Chaetodon ocellatus Chaetodon sedentarius
Spotfin Butterflyfish Reef Butterflyfish
Pomacanthidae Holacanthus ciliaris Pomacanthus arcuatus
Blue Angelfish Gray Angelfish
Mugilidae Mugil cephalus Mugil curema
Striped Mullet White Mullet
Sphyraenidae Sphyraena picudilla Southern Sennet
Uranoscopidae Astroscopus y-graecum Southern Stargazer
Blenniidae Chasmodes bosquianus Hyposoblennius hentz
Striped Blenny Feather Blenny
Gobiidae Gobiosoma bose Naked Goby
Trichiuridae Trichiurus lepturus Atlantic Cutlass fish
Stromateidae Peprilus paru Peppilus triacanthus
Southern Harvestfish Butterfish
Scombridae Scomberomorus maculates Spanish Mackerel
Class: ELASMOBRANCHII (Sharks, Skates)
Family Dasyatidae
Scientific Name Dasyatis Sabina
Common Name Atlantic Stingray
Dasyatis Americana Southern Stingray
Gymnuridae Gymnura micrura Smooth Butterfly Ray
Rajidae Raja eglanteria Clearnose Skate
Sphyrnidae Sphyrna tiburo Bonnet Head Shark
Carcharhinidae Negaprion brevirostris Carcharhimus plumbeus
Lemon Shark Sandbar Shark
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Introduced Non-native Aquatic Species in Florida
Group Species Common Name
Crustaceans-Shrimp Penaeus monodon Asian Tiger Shrimp
Bivalves Perna viridis Green Mussel
Fishes Pterois volitans Lionfish
Mammals Myocastor coypus Nutria
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Useful Websites
Visit these website for good information/definitions on water chemistry topics http://wow.nrri.umn.edu/wow/under/parameters/temperature.html http://waterontheweb.org
Visit this website for lots of information on invasive and nonnative species http://myfwc.com/wildlifehabitats/nonnatives/
Visit this web site for information on animals, plants, water quality, and watersheds http://www.chesapeakbay.net/baybio.htm
Links to information on Oyster Restoration https://www.flseagrant.org/news/2013/09/indian-river-oyster/ Oyster Restoration Workgroup : http://www.oyster-restoration.org http://www.habitat.noaa.gov/pdf/Oyster_Habitat_Restoration_Monitoring_and_Assessment_Handbook.pdf
The Academy of Natural Sciences - Research - Patrick Center - Research Programs http://www.acnatsci.org/research
Florida Sea Grant website https://www.flseagrant.org
Website for Volunteer Estuary Monitoring http://water.epa.gov/index.cfm
EPA Watershed Information http://www.epa.gov/owow/watershed
Conchologists of America- Conch-Net Home Page http://www.conchologistsofamerica.org
Earthguide - Earth Science Educational Resources http://earthguide.ucsd.edu
Center for Watershed Protection http://www.cwp.org/
Marine Protected Areas - News http://depts.washington.edu/mpanews/
Wetland Breaking News http://www.aswm.org/wbn
Weather http://weather.noaa.gov/weather/FL_cc_us.html
Society of Wetland Scientists-Wetland related jobs http://www.sws.org
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US Geological Survey www.usgs.gov
EPA Wetland Fact Sheets http://www.epa.gov/owow/wetlands/facts/contents.html
EPA's Most Frequently Asked Wetland Questions http://www.ehso.com/wetlands_information.htm
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Useful Books on Coastal Wetlands
Aja, D. 1996. A Citizens Guide to Coastal Watershed Survey. Maine Department of Environmental Protection, Maine. 77 pp.
Bahr, L.M, and W.P. Lanier. 1981. The ecology of intertidal oyster reefs of the South Atlantic coast: a
community profile. U.S. Fish and Wildlife Service, Office of Biological Services, Washington, D.C. FWS/OBS-81/15. 105 pp.
Braccia, A. and Batzer, D.P. 2001. Invertebrates Associated with Woody Debris in a Southeastern U.S. Forested
Floodplain Wetland. Wetlands. Vol. 21, No. 1. pp. 18-31 Carpenter, K.E. (ed.). 2002. The living marine resources of the western Central Atlantic. Vol. 1: Introduction,
mollusks, crustaceans, hagfishes, sharks, batoid fishes, and chimaeras. FAO species Identification Guide for Fishery Purposes and American Society of Ichthyologists and Herpetologists Special Publication. No. 5. Rome, Food and Agriculture Organization of the United Nations. pp. 1-600.
Carpenter, K.E. (ed.). 2002. The living marine resources of the western Central Atlantic. Vol. 2: The Living Marine Resources of the Western Central Atlantic. Volume 2: Bony fishes part 1 (Acipenseridae to Grammatidae). FAO species Identification Guide for Fishery Purposes and American Society of Ichthyologists and Herpetologists Special Publication. No. 5. Rome, Food and Agriculture Organization of the United Nations. pp. 601-1374.
Carpenter, K.E. (ed.). 2002. The living marine resources of the western Central Atlantic. Vol. 3: Bony Fishes part 2 (Opistognathidae to molidae), sea turtles and marine mammals. FAO species Identification Guide for Fishery Purposes and American Society of Ichthyologists and Herpetologists Special Publication. No. 5. Rome, Food and Agriculture Organization of the United Nations. pp. 1375-2127.
Coulombe, D. 1992. The Seaside Naturalist. Fireside Publishing, New York, NY, 246 pp. Fischer, W. 1978. FAO species identification sheets for fishery purposes. Western Central Atlantic (fishing area
31). Vols. 1-7. Gilligan, M. 1989. An illustrated field guide to the fishes of Gray’s Reef National Marine Sanctuary. NOAA
Technical Memorandum. Washington, D.C., 77pp. Heard, R.W. 1982. Guide to Common Tidal Marsh Invertebrates of the Northeastern Gulf of Mexico.
Mississippi-Alabama Sea Grant Consortium. Reinbold Lithographing and Printing Co., Booneville, MS, 81pp.
Kaplan, E.H. 1988. Peterson Field Guides, Southeastern and Caribbean Seashores.Houghton Mifflin Co., New
York, NY, 425 pp. Miner, R. 1950. Field Book of Seashore Life. Van Rees Press, New York, NY. 888 pp. Mitchell, M. and Stapp, W. 1992. Field Manual for Water Quality Monitoring: An Environmental Education
Program for Schools. Thomson-Shore Printers, Dexter, Michigan. 240 pp.
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Niesen, T.M. 1982. The Marine Biology Coloring Book. Harper Resource, New York, NY,115 pp. Olsen, M. Georgia’s Wetland Treasures. Georgia Department of Natural Resources, Coastal Resources Division
and U.S. Environmental Protection Agency publication, 218 pp. Pearce, M. 1999. Seashore Life Illustrations. Dover Publications, Mineola, NY. 32 pp. Robbins, C. R., and G.C. Ray. 1986. A Field Guide to Atlantic Coast Fishes. Houghton Mifflin Company,
Boston, MA., 354pp. Ruppert, E.E. and R.S. Fox. 1988. Seashore Animals of the Southeast. University of South Carolina Press,
Columbia, S.C., 428 pp. Stancioff, E. 1996. Clean Water: A Guide To Water Quality Monitoring. Maine/New Hampshire Sea Grant
Marine Advisory Program & University of Maine Cooperative Extension, Orono, ME. 73 pp. Whitney, E., & Means, D. (2004). Priceless Florida: Natural ecosystems and native species. Pineapple Press,
Sarasota, FL. 424 pp. Witherington, B., & Witherington, D. (2007). Florida's living beaches: A guide for the curious beachcomber.
Pineapple Press, Sarasota, FL. 326 pp.
A fishery biologist with the U.S. Environmental Protection Agency examines marsh grass for the presence of marine life in the Florida Panhandle, ca. 1972. (Photo credit: Bill Shroat, EPA)
Science Serving Florida’s CoastPO Box 110400
Gainesville, FL 32611-0400(352) 392-5870
www.flseagrant.org
