Annex 3 MONITORING SEABIRDS IN THE BCLME: DATA …

Top Predators of the Benguela System 1

J. Kemper1

with contributions from:

J. Braby2, B.M. Dyer3, J. James4, R. Jones4, K. Ludynia5, R. Mullers6,J-P. Roux4, L.G. Underhill2 and A.C. Wolfaardt2,7

1 African Penguin Conservation Project, PO Box 583, Lüderitz, Namibia2 Avian Demography Unit, University of Cape Town, Rondebosch 7701, South Africa

3 Department of Environmental Affairs and Tourism, Marine and Coastal Management, Private Bag X2,Rogge Bay 8012, South Africa

4 Ministry of Fisheries and Marine Resources, P.O. Box 394, Lüderitz, Namibia5 Forschungs- und Technologiezentrum Westküste (FTZ) Universität Kiel, Hafentörn 1, 25761 Büsum, Germany

6 University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands7 Western Cape Nature Conservation Board, P Bag X5014, Stellenbosch 7600, South Africa

Final Report of the BCLME(Benguela Current Large Marine Ecosystem)

Project on Top Predators as Biological Indicators ofEcosystem Change in the BCLME

edited by S.P. Kirkman

Annex 3

MONITORING SEABIRDS IN THE BCLME:DATA COLLECTION MANUAL

2 Top Predators of the Benguela System


1. Preamble ................................................................................................................................................... 5

2. Working with seabirds ............................................................................................................................... 5

3. General measuring techniques

3.1 Weighing and measuring an egg ...................................................................................................... 5

3.2 Weighing a bird ................................................................................................................................. 6

3.3 Measuring a bird ............................................................................................................................... 7

4. Mark–recapture techniques ....................................................................................................................... 8

5. Monitoring non-breeding seabirds ............................................................................................................. 8

6. Monitoring seabirds at sea ........................................................................................................................ 8

7. Measuring breeding synchrony ................................................................................................................. 8

8. Checking for pollutants ............................................................................................................................. 9

9. Monitoring disease .................................................................................................................................... 9

10. Measuring daily energy expenditure of chicks and adults, using Doubly-Labelled Water ........................ 9

11. Species-specific monitoring and research techniques

11.1 African Penguin (Spheniscus demersus) ...................................................................................... 10

11.2 Leach’s Storm Petrel (Oceanodroma leucorhoa) .......................................................................... 16

11.3 Great White Pelican (Pelecanus onocrotalus) .............................................................................. 17

11.4 Cape Gannet (Morus capensis) .................................................................................................... 17

11.5 White-breasted Cormorant (Phalacrocorax lucidus) ..................................................................... 19

11.6 Cape Cormorant (Phalacrocorax capensis) .................................................................................. 20

11.7 Bank Cormorant (Phalacrocorax neglectus) ................................................................................. 21

11.8 Crowned Cormorant (Phalacrocorax coronatus) ........................................................................... 21

11.9 African Black Oystercatcher (Haematopus moquini) .................................................................... 22

11.10 Kelp Gull (Larus dominicanus vetula) ......................................................................................... 22

11.11 Hartlaub’s Gull (Larus hartlaubii) ................................................................................................ 23

11.12 Swift Tern (Sterna bergii) ............................................................................................................ 24

11.13 Damara Tern (Sterna balaenarum) ............................................................................................. 24

Acknowledgements ....................................................................................................................................... 25

References .................................................................................................................................................... 25

CONTENTS



1. Preamble

The management of seabird populations within the BenguelaCurrent ecosystem is the responsibility of three countries,Angola, Namibia and South Africa. Within each country, thisresponsibility is shared by a range of agencies. In Angola,management is mainly the responsibility of the InstitutoInvestigação Pesqueira, Ministry of Fisheries and Environ-ment. In Namibia, all offshore islands are managed by theMinistry of Fisheries and Marine Resources. Other seabirdbreeding localities are situated within reserves and thereforewithin the jurisdiction of the Namibian Ministry of Environ-ment and Tourism, while other localities fall under the author-ity of the local municipality or town council, or are on privateproperty. In South Africa, seabird management is also ef-fected by a number of agencies. These include Marine andCoastal Management (MCM), South African National Parks(SANP), Cape Nature, various municipalities and privateproperty owners.

The effective management of the Benguela Ecosystem isthe result of intense research and monitoring of the system.Seabirds form an integral component of this system, and astop predators, are useful indicators of the system’s health.Seabird research has included the monitoring of seabirdbreeding colonies, in particular the numbers of birds breed-ing and how this may change over time and with a changingenvironment. Further research on seabirds as indicators ofecosystem health involves the monitoring of seabird breed-ing success and related parameters, diet and movement.Changes detected in the parameters measured may varytemporally (i.e. over days to years) and spatially (i.e. locallyto regionally).

Given the range of agencies involved in monitoring andmanaging seabird populations within the BCLME, as well asthe high turnover of staff experienced at a number of insti-tutions in recent years, this manual aims to provide “hands-on” guidance on a wide range of appropriate field techniques,in a similar manner to the manual developed for theCCAMLR Ecosystem Monitoring Program (CCAMLAR2004). The manual further attempt to ensure that data arecollected in a standardized manner across the region to al-low comparative analyses between regions, localities andyears. This manual should be viewed as a working docu-ment, subject to periodic revision as monitoring techniquesare refined and improved and new technology is developed.

2. Working with seabirds

Working with seabirds requires great care, patience, anacute awareness of ones surroundings, and a basic under-standing of the behaviour of the species being studied. Thepresence of a researcher can significantly disturb seabirds,and apart from potentially biasing research results, may havedetrimental effects on the individual, breeding colony orpopulation in question. When working close to seabird breed-ing colonies, it is essential to always move slowly and cau-tiously and to avoid stepping backwards before checkingwhat is behind. Always keep a close watch on the reactionof birds near you. Always withdraw with as much care as yourapproach. If there is any sign of behaviour indicating distur-bance, such as nesting birds standing/fidgeting at the nestsuggesting that they may abandon the nest, remain motion-less until the birds resettle, or slowly retreat. Keep the maxi-mum possible distance to seabird colonies (e.g. using a tel-escope rather than binoculars to count a colony of seabirdsreduces the risk of disturbance). Breeding birds sometimestemporarily abandon their nests when disturbed, leavingeggs or small chicks exposed. If predators such as gulls are

present in the vicinity, cover exposed nest contents withsome nesting material to make them less detectable (do notdo this if it would cause further disturbance). When handlingindividuals (e.g. for weighing, diet sampling or logger attach-ment) always have the equipment laid out and ready for useto minimize disturbance. Keep handling time to a minimum.

Work calmly while handling birds. Do not speak in a loudvoice and avoid sudden movements. You may also cover thehead of the bird with some cloth to help calm the bird. If youare bitten or scratched by a bird while handling it, do not dropthe bird. The safety and well-being of the bird should be keptat a premium and handling time kept to a minimum.

3. General measuring techniques

3.1 Weighing and measuring an egg

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Vernier, dial or digital callipers (measuring to 0.1 mm)∗∗∗∗∗ Spring balance and bag (measuring to 0.1 g) OR∗∗∗∗∗ Portable electronic balance and suitable container (meas-

uring to 0.1 g)∗∗∗∗∗ Permanent felt-tipped marker pen (optional)

During incubation, an egg loses weight owing to waterevaporating through the pores in the egg shell. The rate ofweight loss is linear and greater in large eggs (with a largersurface) than in small eggs. Egg measurements can there-fore be used to estimate date of the start of incubation of aparticular nest (Underhill & Calf 2005), and similarly to esti-mate the start of incubation dates for populations at a par-ticular locality (Matanyaire et al. 2002).

Weigh the egg inside a light bag suspended from a springbalance. Alternatively, use an electronic balance to weigh theegg. Place the egg into a container or bag to make sure theegg does not roll off the balance. Subtract the weight of thebag or container from the final weight. If a clutch consists ofmore than one egg, and repeat measurements are to betaken, mark each egg at the sharp egg with small verticallines to be able to identify them again subsequently.

Measure and record the maximum length, as well as themaximum breadth of an egg. To accommodate asymmetri-cal eggs, take two or three measurements of maximumbreadth by opening the callipers slightly, turning the egg in-side the callipers and closing the callipers again (Fig. 1a). Forlarge eggs, callipers may be too short; these eggs may haveto be measured from a different angle (Fig. 1b).

Take care not to damage the egg during measuring:measurements should be taken over one’s lap in case the

Figure 1: (a) The recommended method to measure maximumbreadth of an egg using callipers (egg in side view). (b) the alternatemethod used for large eggs (egg in frontal view). Arrows indicateturning direction of egg to get several breadth measurements, b) isthe preferred way to measure all egg diameters


egg drops, avoid turning the egg repeatedly as this disorien-tates the embryo. Replace the egg in the nest as soon aspossible. If the egg is in danger of being predated before theparent returns to the nest, cover it with some nesting mate-rial.

3.2 Weighing a bird

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Spring balance (balance type and accuracy depends on

the species and the accuracy required) or portable elec-tronic balance (preferable for small chicks)

∗∗∗∗∗ Weighing bag / rope∗∗∗∗∗ Automated weighbridge for penguins (optional)

Adult, immature and large chicks of African Penguins arebest weighed by using a piece of soft rope which is fittedaround the penguin’s body under the flippers, where the flip-pers join the body. Make sure the strap is tight enough toprevent the penguin from slipping out. Hold the bird awayfrom ones body and face to prevent personal injury. Theloose end of the strap is attached to the balance. The birdis then suspended above the ground (Fig. 2). Never take areading while the bird is struggling. Turning one’s back to thewind may reduce movement. Release the bird at the weigh-ing station or at the nest if a chick, by lowering the bird gen-tly and removing the strap with the bird facing away. Tare theweight of the strap or bag prior to weighing, or subtract fromthe final reading.

An automated weigh-bridge can be used to remotelyrecord movement and weight of birds entering or leaving acolony. Birds used for such an experiment have subcutane-ous transponders implanted in the neck region. This mustALWAYS be done by a qualified veterinarian. The organiza-tion wishing to conduct such experimentation must have apeer reviewed ethical process followed.

To set up such an apparatus requires the following action:Identify a suitable colony. Block off the colony or constrict anatural pathway in such a way that it forces birds to walk overthe scale. The scale must be sited at a constriction point of

your choice in the path that the birds use to access thecolony. The scale must incorporate a pre-fitted platform withvertical pitons along each edge length. It must be designedin such a way that it allows only one bird to walk at least threesteps before stepping off (40 × 25 cm is adequate). Place thescale about 25 cm above the ground level. This means“building up” the ground to achieve this. Step this slopesteeply on either side of the scale. This slows the approach-ing birds and facilitates a better reading as the bird crossesthe scale. A transponder reader is placed next to the scaleand between the sensors at a height of approx 30 cm. Infra-red sensors are placed on either side of the scale to recordwhether the bird is entering or leaving the colony. The Infra-red beams, transponder reader and the scale are linked toa PC which captures all the data. There is no ready pro-gramme to install that links all the tools up, so it is advisableto find a specialist to set this up. The PC can either be pow-ered via the mains or with adequate 24V batteries. The PCmust be placed in a weatherproof container and the scalerequires regular maintenance to remove accumulated waterand debris.

Adult Cape Gannets are best weighed in a bag. Place acloth or old sock over the head and tape beak closed. A strapis fitted around the chest and wings, trapping the wingsagainst the body. Great care must be taken not to injure thewings. Place the bird in a bag and suspend from a springbalance. Be very careful when handling gannets at all timesas the beak can cause severe injury. Tare the weight of thestrap, cloth/sock and bag prior to weighing or subtract fromthe final reading.

Smaller species, such as cormorants, gulls and terns, aswell as small chicks of larger species, are weighed inside acloth bag of a suitable size with a drawstring to confine thebird inside the bag if necessary (Fig. 3). Any adult bird mustalways have the head covered and wing strapped in thefolded closed position. The bag must be weighed immedi-ately after weighing the bird and the weight of the bag sub-tracted from the recorded measurement. In windy conditions,face away from the wind for more accurate readings.

Figure 2: Weighing an adult African Penguin by suspending it froma spring balance

Figure 3: Weighing a Cape Gannet chick inside a bag


3.3 Measuring a bird

A number of measurements may be taken from a live or deadbird. Only the most fundamental measurements are givenhere. For greater detail and further measurements, see deBeer et al. (2001).

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Stopped wing ruler∗∗∗∗∗ Ruler∗∗∗∗∗ Vernier, dial or digital callipers

3.3.1 Wing length

Slide a stopped wing ruler under the naturally folded wing tothe carpus and press the carpal joint gently but firmly againstthe stop. Make sure that the wing does not spread. Flattenthe wing against the rule by applying gentle pressure on themedian or greater coverts. Straighten the longest primary bystroking the shafts of the primaries, from the base to the tipwith your thumb while pressing firmly against the rule all thetime. Do not pull the tip of the wing. Measure the distancebetween the carpal joint and the tip of the longest primaryfeather (Fig. 4).

3.3.2 Tail length

Slide a ruler under the tail to where the central pair of tailfeathers emerge from the body. Straighten and flatten the tailif necessary, and measure the longest feather (Fig. 5). Meas-ure to the nearest 1 mm. Do not measure from above, as thismay damage the preen gland above the base of the tail.

3.3.3 Culmen length and depth and total head length

Make sure the bird’s head is properly controlled and takegreat care to prevent injury to the bird’s face and eyes. Openthe calipers wide enough so that the calliper gap is widerthan the culmen / total head length of the bird being meas-ured. To measure the culmen, carefully push the outer calli-per along the culmen to the angle in front of the skull (i.e. to

the base of the feathering or base of the skull, then carefullyclose the inner calliper until the point of the culmen justtouches the inside point of the calliper (Fig. 6).

To measure bill depth, move callipers down length untilyou acquire the widest measurement. In penguins, it is takenwhere the beak and the feathers on the forehead meet. Ingulls and terns it is taken between the widest point on thegonys and the upper beak.

To measure total head length, align the outer calliper withthe longest measurement of the back of the skull, near itsbase (Fig. 6). Align the inner calliper as for the culmen meas-urement. Measure to the nearest 0.1 mm. Always recordwhich method was used to measure the culmen.

3.3.4 Tarsus and toe length

Bend the foot to approximately 90 degrees to the tarsus. Witha ruler or callipers measure the distance between the notchof the intertarsal joint and the lower edge of the last completescale before the toes diverge (Fig. 7). Measure to the near-est 0.1 mm. Toe measurements are taken from the joint be-tween the tarsus and the toe to the base of the claw (Fig. 7).

Equipment use and maintenance

♦♦♦♦♦ Prior to sampling, check that the zero value of spring bal-ances and electronic balances is accurate

♦♦♦♦♦ Balances and weighbridges need to be calibrated regu-larly against objects of known weights.

♦♦♦♦♦ Callipers must be kept in good condition and free ofsand. Light lubrication after usage will facilitate the useof the instrument. Prior to each sampling session, checkthat callipers close or register to zero. If not, check forsand grains between the callipers or in the slidingmechanism. Make sure that digital callipers are zeroedbefore each measurement.

♦♦♦♦♦ Keep tally counters free of sand and lightly lubricated.Regularly check working order by making sure all digits“tick over” smoothly. If digits tend to stick, replace thecounter. Zero the tally counter before each count.

Figure 4: Measuring wing length Figure 5: Measuring tail length

Figure 6: Measuring culmen length of a bird to (a) the base of thefeathering, (b) to the base of the skull, (c) total head length Figure 7: Measuring tarsus length (a) and toe length (b)


4. Mark–recapture techniques

Being able to recognize individuals is vital for estimating arange of demographic parameters for a population of sea-birds, such as movement between localities or regions, sur-vival rates, individual breeding success and age at firstbreeding. There are various ways of marking individuals,including dyes such as Rhodamine B, Picric Acid orPorcimark®, while peroxide (used to bleach dark-backedbirds) can be a useful recognition tool. Transponders im-planted under the skin have also been used successfully torecognize individuals (see also section 3.2).

An automatic recognition system for African Penguins iscurrently being developed. This systems uses the uniquechest spot pattern of African Penguins to recognize individu-als, similar to a bar code system. Equipping individuals withtelemetry devices is gaining popularity as technology devel-ops (see Wilson et al. 2002 for a review).

The most common marking technique currently used inseabirds is to ring individuals and to record details of any re-sightings of that individual. Owing to the invasive nature ofringing, birds may only be ringed by a trained, qualified per-son in possession of a valid ringing permit. An in-depth dis-cussion of ringing techniques is outside the scope of thismanual. Persons interested in becoming ringers should con-tact the South African Bird Ringing Unit (SAFRING) at theUniversity of Cape Town, South Africa about training options(see www.aviandemographyunit.org.za) and consult de Beeret al. (2001).

Once fledged, most seabirds are not easily re-caught. Inaddition to stainless steel rings to identify individuals, colourrings can be fitted. Colours or colour combinations can bechosen to identify different cohorts and breeding localities.Check with SAFRING whether the proposed colour combi-nation has been used for the same species elsewhere. Col-our-ringed individuals provide important demographic datawithout having to be re-caught.

The following basic information should be noted for eachre-sighting or recovery record:

• Ring number on stainless steel ring• Colour ring sequence, where applicable, detailing which leg

colour ring is on• Date• Age group (chick / fledgling, immature, adult)• Locality (e.g. name of island); if ringed individual is encoun-

tered along a stretch of shoreline, provide GPS position• Area (e.g. management zone on island)• Activity (e.g. holding site, nest building, breeding – provide

details of nest contents,); for moulting penguins detail stageof moult

• Special remarks (e.g. sex if seen with partner, any injuries,cause of death if known for recoveries).

5. Monitoring non-breeding seabirds

Monthly head counts of non-breeding seabirds at an islandor along a stretch of shoreline provide useful informationabout the seasonal variation in numbers of birds that use thatarea, and helps to identify particularly important roostingsites.

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope on tripod∗∗∗∗∗ Tally counter

Move slowly along the coastline, using binoculars to lookahead for seabirds. Although the majority of birds will befound along the coast, it is important to also scan inlandsections for roosting birds. Only count non-breeding sea-birds; for counting African Black Oystercatchers refer to sec-tion 11.9.1. Be careful not to count birds more than once;those birds that fly over the observer from ahead should becounted while those that fly from behind should not, as theyhave presumably already been counted. To avoid confusion,only count the birds once you have passed them. Record thefollowing information:

• Observer name• Locality• Date• The number of species counted• The number of individuals counted per species• Time that the count started and ended• Low and high tide times• Age class (adults, subadults, immatures, juveniles); this

may only possible for some species, such as Kelp Gulls.

6. Monitoring seabirds at sea

A range of methods for monitoring birds at sea has beenpublished (e.g. Tasker et al. 1984, Haney 1985, Gould andForsell 1989, Ryan and Cooper 1989, Camphuysen andGarthe 2004). Each method makes a number of assumptionsand may be biased in some way. Use the method whichfulfills the most assumptions, is viable and answers the re-search questions posed. Standard methods may be modifiedto suit the research needs, the vessel’s architecture and itsactivities. For a time series of serial counts of seabirds at seato be comparable, it is vital that the same method is usedeach time. Careful recording of the monitoring method isimportant to ensure that the same method is used in futuremonitoring efforts.

7. Measuring breeding synchrony

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod (optional)∗∗∗∗∗ Tally counter

For most seabirds breeding in the BCLME, annual peakcounts of nesting birds at a breeding locality are used as aproxy for breeding population size there. This approach as-sumes that breeding activities are well synchronized. How-ever, the degree of breeding synchrony may differ betweenseabird species, populations, years localities, and even be-tween colonies at a particular locality. Comparing the breed-ing seasonality patterns of several monitored group at abreeding locality gives an indication of the degree of breed-ing synchrony at that locality and its implication for estimat-ing the breeding population at a particular locality (Kemper2006).

The exact method will depend on the species being moni-tored. For species which tend to breed throughout the year,such as African Penguins, identify several (at least five)groups of nesting pairs. The groups may consist of discretebreeding colonies or demarcated study areas within coloniescontaining at least 30 nests at peak breeding. Every twoweeks, count all active nests (i.e. those containing eggs orchicks) in each group throughout the year. Separately record


nests containing eggs and those containing chicks sepa-rately. Count the number of fledglings in each group.

For species with poor nest site, colony or locality fidelity,such as Cape Cormorants and Swift Terns, choose severalcolonies or sub-colonies (depending on their size) to bemonitored as the birds begin to settle. Choose areas wheremonitoring activities are least likely to disturb a colony. If pos-sible, demarcate the area using natural landmarks or bymarking the area before breeding commences to keep trackof the study area being monitored. Every two weeks countthe number of nests containing eggs, the number containingchicks, and the number of free-ranging chicks and fledglings.Where feasible, binoculars or a telescope should be madeuse of.

8. Checking for pollutants

In addition to oil pollution, industrial pollution is becoming anincreasingly important cause of seabird mortality. Discardedfishing line or ropes may lead to entanglement, while theingestion of plastics, fish hooks and other waste may impairdigestive efficiency or cause internal injuries. African Pen-guins, cormorants, gulls and terns often use such pollutantsas nesting material. Cape Gannets ingest fishing hookswhen foraging near fishing vessels, and may regurgitatethose to their chicks. Gulls and Great White Pelicans tend toforage at rubbish dumps and may regurgitate plastic andother waste at their nests.

In order to investigate the degree of industrial pollutionand its potential effect on seabird populations, dedicated nestchecks for pollutants should be undertaken, particularly atbreeding colonies close to major cities, harbours and fishingareas. Check a random sample of nests (at least 100) forpollutants and record the nature and rough amount (as per-centage of total nest material) found in each checked nest.Pollutants may be divided into categories, e.g. plastics, fish-ing line, ropes. This is best done after breeding has beencompleted.

9. Monitoring disease

Seabirds are susceptible to a number of diseases such asavian cholera, Newcastle Disease Virus (NDV), and (poten-tially) avian flu, or may be poisoned en masse by extensivered tide events. Outbreaks of contagious diseases need tobe managed to ensure minimum mortality.

Seabird mortalities should be regularly monitored throughpatrols at breeding localities as well as along the mainlandcoastline. Record all dead seabirds and note cause of deathif possible. If birds are found dying, carefully record symp-toms. Factors such as stress, starvation, human disturbanceor extreme weather conditions can predispose seabirds todisease. It is therefore important to record such factors as:

• Demographic factors: species, age, sex, reproductivestatus, stage of breeding cycle, colony size

• Environmental factors: locality, date, time, weather, humanaccess and presence, predators

If more dead birds than usual are found on a patrol and dis-ease is suspected, immediately inform the relevant authori-ties and liaise with a veterinary pathologist regarding theidentification of pathogens. Freeze freshly dead birds forsubsequent post-mortem examination by a veterinary lab. Ifmore than one carcass is frozen, label carcasses to keep

track of individual case histories. Incinerate all birds knownor thought to have died from a disease to contain the spreadof pathogens. Wear protective clothing, including gloveswhen handling potentially diseased birds, as some diseasesmay also be contagious in humans. Thoroughly wash outerclothing (oilskins), boots and hands after handling ill or deadbirds. Do not discard any dissection gloves or tools with otherrefuse, rather burn these. Do not use oilskins and gumbootsat more than one locality. Ensure all staffed islands havedissection gear, sample jars and protective clothing.

10. Measuring daily energy expenditure of chicks andadults, using Doubly-Labelled Water

This method is based on the determination of the clearancerates of the stable 2H and 18O isotopes in the animal’s bodywater pool, following a pulse dose labelling of Doubly-La-belled Water (DLW), i.e. water with high 2H, and 18O concen-trations. The difference between the 18O and 2H clearancerates is a measure for the rate of CO2 production, i.e. thebird’s level of energy expenditure. It allows the precise meas-urement of energy requirements of chicks to sustain normalgrowth, as well as of parental daily energy budgets and offlight costs.

This method may be used on a range of seabirds. How-ever, a central requirement of the method is that the studyanimal needs to be captured on several occasions. Thiscondition is met by relatively “docile” species such as CapeGannets, which can be readily caught at their nest sites with-out causing undue stress to either the study animal itself norto nearby breeders. This method is not recommended forspecies such as cormorants, which are highly susceptible todisturbance and are difficult to catch.

The kinetics of the 2H and 18O isotopes in the bird’s bodywater pool are assessed from 60 µl blood samples, takenbefore DLW administration (“baseline”), 60 minutes afteradministration (“initial”), and 24 hours thereafter (“final”). Thedose (0.5 g per kg body mass), injected intra-peritoneally, issufficient for 24–48 h measurements. The same mass-specific dose can be applied in adults and in chicks. DLWadministration and blood sampling may only be undertakenby a trained person or a veterinarian and is subject to priorapproval from a relevant authority (e.g. an animal ethics com-mittee or ministry). Blood samples (preferably in triplicate)need to be further evaluated by isotope specialists.

11. Species-specific monitoring and research techniques

Monitoring techniques particular to each seabird speciesbreeding in the BCLME are listed below. Species are listedfollowing the taxonomic sequence used by SAFRING (deBeer et al. 2001).

A comment on counting…

♦♦♦♦♦ Use a tally counter for counting large groups of birds ornests; check it frequently to ensure that it is adding upcorrectly.

♦♦♦♦♦ Make several (two to three) counts of the same group,either by repeating your count or having a second per-son counting

♦♦♦♦♦ Compare counts. If counts differ by more than 10% makeanother count.


11.1 African penguin (Spheniscus demersus)

African Penguins in adult plumage are patterned black aboveand white below, with a characteristic white head stripe andblack breast band. Juvenile African Penguins (betweenfledging and their first post-fledging moult one to two yearslater) are grey to brown above with white under-parts and abroad brown breast-band. During this first (juvenile) moultafter fledging they acquire their adult plumage and becomeindistinguishable from sexually mature adults. Some juvenilebirds undergo a partial head moult at sea (Fig. 8).

While breeding African Penguins frequently opt to remainon their nest and threaten intruders rather than flee, thepotential for nest abandonment does exist. It is thereforeessential that disturbance is kept to a minimum, regardlessof whether a colony or an individual appears to be “relaxed”or not.

11.1.1 Catching African Penguins

African Penguins can be caught by hand, a gannet crook ora sturdy hoop net attached to a short pole, or a noose at-tached to the end of a pole (see sections 11.4.1 and 11.5.2,Figs. 24 and 26) can also be used. Alternatively, groups ofpenguins can be herded into a coral made with moveablelightweight screens, before catching. It is recommended thatpenguins be captured outside the breeding colony wherepossible, not inside it. If the penguin’s escape route is re-stricted (for example if it is inside a burrow) or if it holds itsground when approached, move toward the penguin veryslowly, watching for any signs of the penguin preparing toflee. Take care not to disturb other penguins in the vicinity.Once the penguin is within reach, crouch down and swiftlygrab the penguin firmly behind the head (NOT the neck asthis could cause serious injury to the penguin). After estab-lishing a good grip, move the other hand under the belly tosupport the body before picking it up. Always make sure thatthe beak is pointed away from your face and body. Alterna-tively, if practical, gently pull the penguin towards you by aflipper. Penguins tend to strain in the opposite direction,usually offering an opportunity to get a good grip of the backof the head. Penguins breeding on surface colonies are bestcaught with a crook (section 11.4.1, Figure 24) or a pole witha noose attached (slip the noose over the head and turn thepole, which tightens the noose; it is then possible to pull thebird toward you), to minimize disturbance of neighbouringbreeders. The method requires some practice as the rela-tively short penguin necks makes it difficult to get a good grip

with the crook.Catching African Penguins at a landing stage (beach) is

more difficult than at their nest sites. In the absence of agannet crook, net or herding screen, approach the penguinsfrom the sea side to prevent them escaping. Catch in theprescribed manner, but resist running after them and caus-ing disturbance to other penguins. The procedure should beabandoned if undue disturbance is caused.

11.1.2 Breeding population estimates

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod for inaccessible or easily disturbed

areas∗∗∗∗∗ Tally counter∗∗∗∗∗ Short, smooth plank or pole (optional)

11.1.2.1 Breeding population estimates from active nest counts

The purpose of active nest counts is to estimate the numberof breeding pairs at a particular locality. Active nests aredefined as nests containing eggs or chicks (Kemper et al.2001). Nests containing medium-sized or larger chicks areeasily counted. Nest sites defended by an adult sitting on anempty nest bowl can be difficult to distinguish from nestsbeing incubated or containing small chick. Inspecting nestcontents by physically handling penguins during an activenest count is not recommended for surface nests. For bur-row counts, a long plank or thick pole may be used to slowlyand gently push against the bird’s chest and to gently lift upthe penguin far enough to inspect potential nest contentswithout harming them. Use a torch to see into deep burrowsor inside buildings.

Counts of surface nests should be done from a distance,using either binoculars or a telescope. Nest sites then needto be distinguished from active nests from visual clues. Pen-guins defending an empty site are more likely to stand up orget off the nest when approached and the site does not nec-essarily contain any fresh nesting material, or no nestingmaterial at all (Fig. 9). Penguins guarding eggs or smallchicks tend to sit on the nest more tightly on and lower in thenest bowl than penguins guarding an empty site and theirnests contain fresh nesting material (Fig. 10).

As the chicks get older, they will become visible under theparent guarding them. Chicks are generally constantlyguarded until the age of about 40 days (Seddon and vanHeezik 1993). Thereafter, they may be left alone for variable

Figure 8: African Penguin in juvenile plumage after undergoingpartial head moult

Figure 9: Active nest site: note the lack of fresh nesting material andthe loose stance of the penguin on the site. This penguin is holdingthe site, but is not guarding nest contents


periods of time and may congregate and form groups or“crèches”. All chicks in a crèche must be counted and re-corded. The total is then divided by two to represent thenumber of active nests. For example a group of three chickswill equal two active nests; a group of ten chicks will equalfive active nests. When doing a count of burrows or of pen-guins nesting solitarily or in loose groups, a mark on theground next to the nest or group of nests can be helpful tokeep track of which nests have been counted. The use of acolour chalk mark on rocks is also useful when having tocount birds breeding in such habitats.

Nest counts are the most common census method appliedat African Penguin breeding localities, and the most practi-cal census method in areas which are seldom visited. At lo-calities where it is feasible, particularly where breeding ispoorly synchronized and there are several breeding peaksin a year, monthly counts of active nests would be useful.Annual breeding population estimates using nest countscould be considerably improved by counting discrete colo-nies at a locality separately and by summing the annual peakcounts of the discrete colonies. At localities where it is notfeasible to do monthly nest counts, (e.g. Dassen Island),monitoring of specific sites is recommended in addition to anannual comprehensive count.

11.1.2.2 Breeding population estimates from active nest sitecounts

Not every breeding pair may necessarily breed in a given year.Active nest counts therefore provide an indication of thenumber of pairs breeding at a given time rather than give es-timates of the potential breeding population. Active nest sitecounts have been used to gauge the size of the potentialbreeding population (including pairs not breeding for somereason). Active nest sites are defined as nests containing eggsor chicks, nests containing fresh nesting material or freshguano, or nests defended by at least one adult (Shelton etal. 1982). At a number of localities (particularly in South Af-rica) active nest sites rather than active nests are counted.

A word of caution…

For breeding population estimates to be comparable be-tween localities and years, the method used to estimate abreeding population must be clearly specified during moni-toring activities to avoid potential misinterpretation of timeseries of nest counts.

11.1.3 Population estimates from moult counts

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod∗∗∗∗∗ Tally counter

Counts of moulting African Penguins have been used toestimate the number of penguins attached to a particularlocality or region (Randall et al. 1986, Underhill and Crawford1999). Penguins moult annually to replace worn and dam-aged feathers. The feather-shedding stage lasts approxi-mately 13 days, during which time all its feathers are re-placed. Penguins are confined to land for that period. Justprior to commencing moult, the feathers unlock and startstanding out from the body, giving the individual the appear-ance of a fluffy pillow (Fig. 11). Feathers may be shed andreplaced on the body in any order, but often feathers are firstshed on the swollen flippers and the face and the last feath-

Figure 10: Incubating penguin: note the abundance of nesting ma-terial and tight stance on the nest, with flipper held close to the body

Figure 11: African Penguin at the beginning of the feather-sheddingstage of moult. Note all feathers standing up and first feathers fallingout

ers are usually shed on top of the head and the back of theneck. Dirty penguins, especially those which have been sit-ting on a nest for an extended period, sometimes appear tobe moulting, particularly if some feathers stick out from thebody. It is important to ascertain that a penguin is indeed inmoult and not simply scruffy or hot.

The moult from juvenile into adult plumage must be re-corded separately from adult moult. Care needs to be takento distinguish age groups in penguins which have nearlycompleted moult. To distinguish between adults and juve-niles check the colour of the feathers being replaced on thesides of the head. Adult penguins have a white band of feath-ers (Fig. 12) while those in juvenile individuals are brown(Fig. 13). However, this may be difficult to detect in those ju-veniles which have previously undergone a partial headmoult (Fig. 14). In addition to moulting juvenile penguins,separately record the number of juvenile penguins NOT un-dergoing moult. This gives an indication of the seasonalityof juvenile penguins returning to land to moult.

Moult counts should be done at two-weekly intervalswhere practical (Randall et al. 1986). This may, however, notbe feasible at remote localities. For these localities, at leastone but preferably several counts around the adult and ju-venile moult peaks should be done. The timing of peak moultis locality-specific (Table 1), and will vary slightly betweenyears. Moult appears to be less synchronized at localities inNamibia than at localities in South Africa, and more countsaround the peak may be required to improve accuracy of theestimate.


11.1.4 Breeding success (and associated parameters)

Equipment needed:

∗∗∗∗∗ Notebook / nest forms and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Balance(s)∗∗∗∗∗ Markers for staking out nests (optional)∗∗∗∗∗ Banding equipment (optional)∗∗∗∗∗ Other marking equipment such as dye or wool (optional)∗∗∗∗∗ Short, smooth plank or pole (optional)

Egg-laying in African Penguins is rarely observed and onlynest sites where nest-building or mating activities are notedor nests which contain eggs which appear to have been re-cently laid should be chosen. Never choose nests, wherenest checks could cause disturbance of neighbouring nests.A minimum sample of 30 nests should be monitored at alocality at any given time. To get an indication of parentalcondition, weigh both parents at the beginning of the nest-ing attempt to assess their condition. If possible, weigh theparents again shortly before the chick(s) fledge(s). If this islikely to cause nest abandonment, assess the condition ofparents from visual clues at the beginning and end of thebreeding attempt.

Nest contents need to be recorded regularly. The nestcheck schedule will depend on the type of informationneeded while minimizing potential disturbance. Weekly nestchecks are adequate for establishing basic breeding successin terms fledglings produced per nest or per pair. More fre-

Figure 13: Moulting juvenile individual. Note brown feathers beingreplaced by characteristic adult white head stripe

Figure 12: Moulting adult individual. Note white feathers being re-placed on the white head stripe

Table 1: Timing of juvenile and adult moult peaks for localities for which the timing is known. Timing is expressed as moult peak month orpeak half month. FH = first half, SH = second half

Breeding locality Position Juvenile moult Adult moult

Mercury Island 25°43'S 14°50'E FH January SH December, FH MayIchaboe Island 26°17'S 14°56'E FH January SH December, FH MayHalifax Island 26°37’S 15°04'E FH January FH January, SH AprilPossession Island 27°01'S 15°12'E January SH December, FH MayBird Island (Lambert’s Bay) 33°05'S 18°18'E November SH October, FH NovemberDassen Island 33°25'S 18°05'E December DecemberRobben Island 33°48'S 18°22'E FH December, March FH DecemberThe Boulders 34°11'S 18°27'E FH November FH DecemberStony Point 34°22'S 18°54'E FH November SH November, FH DecemberDyer Island 34°41'S 19°25'E November, March NovemberSt Croix Island 33°47'S 25°46'E FH December SH November, FH December

quent nest checks might be required if questions such aschick growth, age at fledging, cause of nest content mortal-ity are to be addressed.

Record nest contents during each visit, with an indicationof chick age. Categorize chick ages as follows (also seeSeddon and van Heezik 1993 for details):

• P0 – Hatchling (c. 1 day old; eyes shut)• P1 – Small downy chick (c. 2–10 days; covered by adult,

crawling and raising head)• P2 – Medium downy chick (c. 11– 20 days; not fully cov-

ered by adult, may sit next to adult)• Large downy chick (c. 21–35 days)• ¼-shed (starting to lose down)• ½-shed (half of down replaced by feathers)• ¾-shed (nearly fully feathered)• Blue (fully feathered)

If necessary, use a pole to gently push against or underneaththe adult’s breast and gently lift the penguin far enough toinspect nest contents. However, it is essential to keep dis-turbance and contact to a minimum to prevent the adult fromabandoning the nest. Penguins nesting in sheltered sites,such as in burrows or under bushes are less likely to fleeduring a nest inspection than those nesting on the surface.If the parent flees the nest, leaving eggs or small chicks ex-posed, cover these with some nesting material to preventgulls from predating nest contents, or guard the chicks ashort distance away from the nest, until the parent returns.

To compare differences in breeding success betweennest types, note key characteristics of each nest relevant to


the aim of the study. Attributes could include but are not re-stricted to:

• Nest type (e.g. burrow versus surface nest)• Colony size• Nest position (edge versus centre)• Susceptibility to flooding• Susceptibility to predation

For estimates of chick growth, chicks need to be weighedregularly. Start weighing chicks only after the hatchedchick(s) eyes are fully open. Chicks can be weighed every5 days until they have lost all their down. Thereafter they canbe weighed every 10–14 days until fledging. Weigh chicks tothe nearest 10 g if below 1 kg, and to the nearest 25 g onceabove 1 kg. Specify type and accuracy of the balances used.All chicks should be weighed before fledging, once fullyfeathered.

Chicks may be defended by an adult (or in some casesboth parents). Avoid checking nests or weighing chicks whileparents change shifts, undergo their greeting ritual and whilechicks are being fed. As monitored chicks become moreindependent and show signs of spending time in crèches orin neighbouring burrows, they should be temporarily marked.One option is to mark chicks with brightly coloured dye on thelighter body parts (e.g. on the chest or the underside of theflippers), using different dye patterns and colours to distin-guish between marked chicks. Do not mark the individualexcessively. Dye-marking may have to be repeated asdowny chicks become feathered.

Alternatively, penguin chicks may be temporarily bandedwith a piece of wool fitted loosely around the flipper near theshoulder. The wool band should be removed before the chickfledges; it may be replaced by a permanent penguin flipperband at that time. The wool should be thin enough to tearwithout injuring the chick if it gets hooked in vegetation. Thewool should also be able to break up easily in sea water, incase the chick fledges before the wool can be removed. Sib-lings may be wool-banded on different flippers to tell themapart. Different-coloured wool can be used to mark chicksfrom different nests, particularly if monitored nests are closeto each other and chicks from several monitored nests arelikely to crèche.

If breeding success is to be measured per pair per year,the breeding efforts of individual pairs must be monitored(Crawford et al. 2006). To ensure the identification of indi-vidual pairs over several breeding attempts, band each part-ner and mark the nest. Sexes are generally difficult to tellapart, but the smaller penguin of a pair (usually with a slightlysmaller, more slender beak) will most likely be the female.Possible sex should be recorded for each banded individual.If nest success rather than pair success is monitored, it is notnecessary to follow the same individuals.

11.1.5 Re-sightings of banded individuals

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope / tripod (optional)

The asymmetrical steel bands used for banding individualAfrican Penguins have been designed to be fitted on the leftflipper. However, some bands have been designed to fit onthe right flipper and a few bands have been mistakenly fit-ted on the right flipper. Depending on the series, the numberwill consist of one letter followed by either four or five digits.The number on the band can usually be read with the aid of

binoculars or a telescope. Re-sighting work may be doneopportunistically while carrying out other activities, or throughregular, scheduled sessions.

Scan groups of penguins at landing beaches or coloniesfrom different angles using binoculars or a telescope. Do notcause disturbance to the penguins while moving.

Once a banded penguin has been located, read thenumber on the flipper band. Make sure that the number iscorrect before recording it. In some cases, parts of thenumber may be obscured, for example in moulting or dirtypenguins. Only record complete numbers and those that youare sure are correct. If necessary, change your position tosee the number from a different angle. Record any bandseen to have caused feather wear, injury or death.

11.1.6 Diet sampling using a stomach flushing technique

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Oilskin pants and gumboots∗∗∗∗∗ Rubber gloves∗∗∗∗∗ 25 litre bucket∗∗∗∗∗ 500 ml beaker∗∗∗∗∗ Tablespoon∗∗∗∗∗ Balance∗∗∗∗∗ 500 ml funnel∗∗∗∗∗ 6 mm enema-tube∗∗∗∗∗ Lukewarm water (preferably fresh)∗∗∗∗∗ Sieve with mesh size of 1 mm∗∗∗∗∗Plastic sample bags (freezer bags) or 1 litre plastic sam-

ple jars∗∗∗∗∗ Permanent marker pen∗∗∗∗∗ Waterproof labels and pencil∗∗∗∗∗ Banding equipment (optional)

Diet samples from penguins are collected by inducing thebird to regurgitate its stomach contents (CCAMLR 2004)using a water offloading technique. This procedure can bedone by one person, although it is more practical with twopeople. Only trained people should perform this technique.If there is ANY risk of injuring the bird during the procedure,abandon the process. Have the equipment ready before han-dling birds.

Catch individuals as they arrives from the sea, never atthe nest. After recording the weight of the bird, place it firmlybetween the knees, with the penguin body suspended be-

Figure 14: Juvenile moult of an individual which had previouslyundergone a partial head moult. Note the few brown feathers in thewhite head stripe and below the black chest band


tween the handler’s knees and the flippers resting on han-dler’s lap. Only commence tubing the bird when it hasceased struggling. The bird will usually open its beak to bitewhen shown a hand or finger. When it does this, quickly in-sert the fore and middle finger sideways up to the gape (dothis forcibly if the bird is not obliging). Turn fingers sidewaysso that one can open and close the fingers inside the beak.You are also now able to control the beak by gripping it withyour thumb and ring finger. Open the beak by spreading yourfore and middle finger. Look down the throat to identify thewindpipe (the orifice that frequently opens and closes behindthe tongue). NEVER put the tube into that opening. It is quitedifficult to force the tube down the windpipe and usuallymeets with resistance and tends to cause bleeding. NEVERcontinue working with a bird if this happens; treat the bird andrelease it.

Gently stretch the neck of the penguin to facilitate the pas-sage of the wetted tube which is connected tightly to a 500 mlfunnel at the other end. In the case of resistance to the tube’spassage, gently and slowly remove the tube and try again.Never force the tube. If resistance persists, rather reject thepenguin and look for another individual. Always double-check that the tube is inserted correctly before proceeding.

Once c. 20–25 cm of tube has been inserted (the approxi-mate distance between the base of the lower beak and thelower end of the sternum) or until resistance is felt, pourwater into the funnel. Raise the funnel directly above thebirds head. If water is not draining, carefully nudge the tubeup and down the throat until you note the water draining.Water blockages are usually caused by large food chunks inthe stomach or by the tube getting pinched in the throat orthe stomach. While water is draining from the funnel one cancheck to see if the bird has food or not. Lower the funnelbelow the birds head – if green liquid feeds back up the tubethe bird is empty and you can remove the pipe and releasethe bird – if brown or pink water or particles are present thanthere is a sample.

Allow water to drain either until the funnel empties, thebird begins to make gargling noises or when water runs fromthe beak. Stop sooner if the abdomen feels very distended.At this point, crimp the tube and gently but swiftly remove it.Stand up and while still holding the bird between the knees

invert the bird over a sieve or bucket (Fig. 15). With your freehand massage the throat toward the head while simultane-ously applying pressure to the lower stomach region of thebird with your knees. Do not squeeze excessively, as thismight rupture the stomach. Continue massaging until you feelthat most of the sample has been retrieved. If the bird hastrouble regurgitating because the food is lumpy, let the pen-guin rest in an enclosure for up to half an hour to allow thefood to mix with the water and to soften up. Sometimes largeprey are hard to dislodge and may have to be carefully pulledfrom the throat. With large, soft samples food particles gettrapped in the throat and tongue. The penguin may be in-verted once more if necessary, but not more often to avoidexcessive stress to the bird. Wash the mouth and tongue withwater from the beaker before releasing the bird.

Samples must be carefully removed from the sieve orbucket with a spoon. Do not break up whole prey items tofacilitate identification and measurements. Sample must beput into a sample bag (or jar) and labelled with a marker pen(species, date, locality, sample number, and, if applicable,band number of bird). An additional waterproof label with thesame information written in pencil must be added inside thebag, in case the labelling on the bag rubs off during freez-ing. The sample must be frozen if it cannot be processed im-mediately. The bird should not be handled for more than 10minutes.

If there is a banding and re-sighting programme at thelocality where the diet sample is taken, you may band the in-dividual (while it is already being handled). After recordingbanding details, release the penguin close to where it wascaught.

At staffed localities, diet sampling should be undertakenregularly to detect seasonal changes in diet. Weekly samplesfrom eight individuals are sufficient to yield statistically reli-able results. At remote localities, ad hoc samples may stillbe valuable, but results need to be interpreted with caution.

11.1.7 GPS logger deployment

GPS loggers are used to study foraging ecology (includingat sea behaviour and movement at sea) and provide infor-mation on foraging areas used by African Penguins. Thedevice may include temperature and pressure sensors. GPSloggers have an excellent spatial resolution (10–20 m) andmay provide highly detailed information, with up to one read-ing per second. The disadvantage of a GPS loggers is its lim-ited battery life of a few days. GPS loggers are thereforemostly used to investigate foraging ecology in breeding pen-guins, because the device is likely to be recovered within afew days.

An extensive manual is supplied by the manufacturer ofthe device, explaining the functions and programming of thedevice. Read the instructions carefully before using the de-vice. Perform a test run using the logger “on land” beforehandling birds in order to get accustomed to the preparing,programming and data downloading. Select the settings tosuit your exact research needs.

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ GPS logger – started and closed properly∗∗∗∗∗ TESA tape – black for penguins∗∗∗∗∗ Plastic logger outline (stencil)∗∗∗∗∗ Scissors∗∗∗∗∗ Scalpel or knife∗∗∗∗∗ Balance∗∗∗∗∗ Rubber glue (Pattex)

Figure 15: Inverting an African Penguin over a sieve to collectinga diet sample


Choose nests with medium downy chicks, since at that chick ageone parent stays at the nest during the day. Chances that thebird leaving to sea returns in the evening in order to feed andswap nest duties with the other parent are high and make recov-ery of the logger easy.

Monitor nest attendance in the study colony beforehand. Ifbirds leave early in the morning or during night hours, the attach-ment of the logger can take place in the late evening hours whenthe partner has returned from sea and both parents are presentat the nest site. Make sure to attach the logger to the partnerwhich has spent the day at the nest and will leave the same nightor next morning to sea. This partner is usually dry and dirty whilethe one that has just returned from sea will be clean and possi-bly still wet. Deploying the bird after the partner has returned hasthe advantage that the chick(s) will be protected by one parent,while the other parent has a logger attached.

Attaching a GPS logger to a penguin is easiest with two peo-ple (Wilson et al. 1997). One person catches and holds the birdwhile the other person prepares the deployment and attachesthe logger to the bird. Taking care not to disturb the rest of thecolony, slowly approach the nest, catch the bird to be equippedand take it out of the colony. Cover its eyes to reduce stress.Weigh the bird before attaching the logger.

After weighing, the handler holds the bird calmly on the lap,presenting the bird’s back straight towards the person fitting thelogger. Stick the plastic logger outline template with some stripsof tape to the birds lower back. Make sure it is positioned straightand centred on the lower back (Fig. 16).

Take a piece of tape about 10–15 cm long and hold it be-tween your fingers with the sticky side up. Use the scalpel orknife to lift some feathers in the upper part of the plastic outlineand place the tape with its sticky side facing upwards underneaththe feathers so that they stick onto the tape, leaving the remain-ing ends of the tape on the left and right side hanging down (Fig.17). Repeat the procedure with the next layer of feathers down-wards from the strip of tape just applied. When reaching thelower end of the outline, you should have placed 4–5 strips oftape (Fig. 18).

Place the logger onto the feathers which are sticking to thetape and fasten the tape across the logger (Fig. 19). Always startwith the right end of the tape, wrapping it tightly onto the logger,without leaving folds in the tape, then stick the left end of thesame strip to the logger. Always starting with the right end willhelp when undoing the tape during device recovery, especiallywhen working in the dark or with a nervous bird. The tape at thenarrow top end of the logger might need to be cut to preventfolds. Once all tape ends have been firmly secured, the plasticoutline can be taken off. This is done by pulling it gently down-wards over the logger to prevent feather damage (Fig. 20). Toreduce gaps between logger, tape and feathers where the birdcould tug, secure another strip of tape around the border of thelogger (Fig. 21), except for the bottom part of the logger wherethe pressure sensor is located. To avoid folds, some small cuts

Figure 16: Attaching the logger outlinetemplate cut out of strong plastic onto thebirds back

Figure 17: Applying the first strip of tape Figure 18: Reaching the bottom end of thetemplate

Figure 19: Wrapping the tape tightly across the logger

Figure 20: Removing the template

Figure 21: Placing tape around the logger

Figure 22: Sticking loose tape ends together with glue


in the tape at the upper area of the logger may be necessary.To prevent the tape from unravelling in the water, a thin layerof rubber glue may be used to secure the loose ends (Fig.22). Care must be taken not to get glue on any feathers. Theglue should dry immediately. The penguin must then be car-ried back to its nest without disturbing surrounding birds (Fig.23).

The nest site should be monitored regularly during the fol-lowing evening. Remove the logger as soon as the penguinreturns to its nest, as it may try to remove it itself! After catch-ing the bird and taking it from the colony, the handler shouldagain hold the bird, while the other person peels each layerof tape from the logger. Once the logger is taken off, the lay-ers of tape still sticking to the feathers can be easily removedby pulling the tape gently downwards while holding thefeather bases down with a finger, thus preventing the feath-ers from pulling out. Once the logger and tape have beenremoved, release the penguin at or near its nest.

11.1.8 PTT deployment

PTTs (Platform Transmitter Terminal) function for six to eightweeks, sometimes longer, depending on the preset dutycycle and the age of the batteries. These devices are use-ful for longer-term studies than those using GPS loggers, forexample for examining the foraging ecology of non-breedingindividuals or an entire chick-rearing attempt. The position ofthe penguin is recorded from a signal transmitted by thePTT’s antenna to satellites passing overhead. No signal willbe received if no satellite is in the area during transmissionor when the bird is diving. At least two messages need to betransmitted by the PTT during a satellite’s overpass to pro-vide a usable position. The data is downloaded from thesatellite to the Argos data collection system in France. Datacan therefore be obtained while the bird carrying the PTT isstill at sea. Positions are less accurate (up to 150 m, butpossibly only to the nearest 1 km) than those obtained froma GPS logger. For the exact functioning of the device andhow it is programmed, refer to the manual supplied by themanufacturer. Test the PTT before application to ensure thatit is transmitting properly.

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ PTT device – ready to be deployed∗∗∗∗∗ Balance∗∗∗∗∗ Bag or sock∗∗∗∗∗ Plastic device outline template∗∗∗∗∗ TESA tape∗∗∗∗∗ Scissors (for deployment of less than one week )∗∗∗∗∗ Scalpel or knife (for deployment of less than one week )∗∗∗∗∗ Rubber glue (for deployment of less than one week )∗∗∗∗∗ Velcro (for deployment of more than one week)

∗∗∗∗∗ Araldite epoxy (adhesive and catalyst) (for deployment ofmore than one week)

∗∗∗∗∗ Cloth with acetone∗∗∗∗∗ Flat knife or tongue depressor (for deployment of more

than one week)∗∗∗∗∗ Latex gloves∗∗∗∗∗ Mixing board

If the PTT device is to be deployed for a relatively short time,attach the device in the same manner as a GPS logger (seesection 11.1.7). If the device is to be deployed for more thanone week, Araldite adhesive is used to attach the device toensure that it does not drop off. The PTT is held to the lowerback of the penguin by a strip of Velcro (the size of the PTT)glued to the underside of the device and a correspondingstrip glued onto the back of the penguin. Bear in mind thatthe Araldite sets quicker at higher ambient temperatures.

One person catches, weighs and holds the bird while cov-ering its eyes with a bag or sock, while the other person pre-pares the deployment. Hold the penguin in the same man-ner as outlined for GPS logger deployment (section 11.1.7).Use acetone applied to a cloth to remove natural oils fromthe work surface on the birds back. Tape a template of theshape of the PTT to the mid-lower back of the penguin (sameposition as for GPS loggers (section 11.1.7, Fig. 23). Mix thecomponents of the glue in equal amounts on a board withseparate spatulas. Apply the mixed resin directly onto thefeathers inside the outline template with a spatula or similarobject. Take utmost care to ensure that excess resin doesnot run onto feathers outside the template and not to touchany other parts of the bird. Do not work the resin deep intothe feathers, because the mixture gets hot while setting andmay burn the skin if there is contact. Carefully place theVelcro onto the resin, making sure that it fits cleanly and thatthere is no overspill. Only remove the template once the gluehas dried. Attach the PTT ensuring that it fits squarely ontothe Velcro strip. Do not forget to remove the magnet on thePTT to activate it. Release the bird and observe how the birdmoves with the attached PTT.

To recover the device, remove it by detaching the Velcrostrips from each other. Leave the corresponding Velcro stripglued to the penguin’s back to fall off by itself. Do not attemptto remove the strip as this will cause feather damage and willsubsequently compromise the bird’s insulation.

New PTT models are constantly being developed. Theseare designed to last longer and provide additional informa-tion (e.g. details on water temperature). Because adhesivestends to break up in salt water after about eight weeks, suchloggers should be deployed with a harness system.

11.2 Leach’s Storm Petrel (Oceanodroma leucorhoa)

Although a common summer visitor to south-westernAfrica, little is known about the breeding status of thesecretive Leach’s Storm Petrel in southern Africa. In SouthAfrica, breeding has so far been recorded at three islands,where they mainly breed in rock crevices or stone walls.Leach’s Storm Petrels are seldom seen at their breedingislands, because they feed at sea during the day and onlyvisit their burrows at night.

11.2.1 Breeding population estimates and breeding success

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Recording of Leach’s Storm Petrel call∗∗∗∗∗ Tape recorder or MP3 player with speakers

Figure 23: Penguin with GPS logger with partner at the nest site


∗∗∗∗∗ GPS∗∗∗∗∗ Torch∗∗∗∗∗ Remote camera system (optional)

Because of their secretive nature, breeding Leach’s StormPetrels are difficult to count. On a moonless night, play arecording of their call in areas where they are likely to breed.If Leach’s Storm Petrels are in the vicinity, they tend to im-mediately respond to the call (Whittington et al. 1999). Fromthe number of calls, estimate the population size and locatethe position of potential nests. Record the GPS position ofnests. Using a torch, confirm the presence of a nest. Do notunduly disturb breeding individuals.

Once a nest has been located, it can be monitored usinga remote camera system, similar to a system designed to ex-amine the inside of sewerage pipes. The camera, attachedto the end of a flexible cable is manoeuvred with the cableand a thin cord attached to the head of the camera. This isthe least intrusive method to follow the success of individualnests. Repeated nest checks by sticking an arm into theburrow and feeling nest contents are likely to cause exces-sive disturbance and is not recommended.

11.3 Great White Pelican (Pelecanus onocrotalus)

Adult Great White Pelicans have a white body; breeders aredistinguished from non-breeders by a swollen, knob-like fore-head and duller bare parts. The body of juvenile pelicans isa dull brown with a white rump. Great White Pelicans arecolonial, high density ground nesters. The exact colonylocation may vary between years. Breeding pelicans arehighly susceptible to disturbance and should not be ap-proached closely during the breeding season. Great WhitePelicans may prey on the chicks of other seabirds, includingAfrican Penguin, Cape Cormorant and Kelp Gull.

11.3.1 Breeding population estimates

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod∗∗∗∗∗ Locality map∗∗∗∗∗ Aerial photograph (optional)

Count pelican colonies monthly from a distance with the helpof binoculars or a telescope. If possible, count nests from anelevated vantage point. Record colony position and extenton a map of the locality. If accurate ground counts are notpossible, use an aerial photograph of the colony, taken dur-ing the breeding peak. Individual nests are easily distin-guished on aerial photos, and cannot be confused with thenests of other seabird species.

11.3.2 Ringing fledglings

Equipment needed:

∗∗∗∗∗ Herding screens (aluminium frames covered with netting,2 m x 1 m)

∗∗∗∗∗ Portable fencing∗∗∗∗∗ Ringing equipment∗∗∗∗∗ Balance (optional)∗∗∗∗∗ Ruler (optional)

Pelicans may be ringed just prior to fledging, just before theyare able to fly. This can only be done by rounding up a group

of fledglings with a team of field workers. Do not choose anexcessively hot day, to avoid undue heat stress of capturedchicks. Discuss the best way of how to approach the colony,with the aim of herding as many pelican chicks into a cap-ture pen as possible. The exact strategy will depend largelyon the terrain of the colony. Surround a group of fledglingswith herding screens, herd them a short distance away fromthe colony and corner them by forming an enclosure with theherding screens. Alternatively herd the group into an enclo-sure made from portable fencing which is set up beforehand.Ensure that the enclosure is sturdy enough to withstand agroup of chicks pushing against it.

Remove fledglings one by one from the enclosure for ring-ing, weighing and measuring. Release them near the colony(without disturbing the rest of the colony). Always keep aneye on the captive birds to monitor any risk of overheatingor suffocation. Return visibly stressed individuals to thecolony straightaway if necessary.

11.4 Cape Gannet (Morus capensis)

The Cape Gannet is endemic to southern Africa, where theentire world population is distributed over only six breedinglocalities. The adult plumage is mainly white, with a paleyellow head. Primaries, secondaries, primary coverts and tailare black. Juvenile Cape Gannets are dark grey-brown withwhite speckles, with varying amounts of white on the neck.The juvenile plumage is gradually replaced by white, start-ing on the lower body and finishing on the wings. Individu-als are generally more docile than African Penguins andtherefore easier to handle. However, always ensure that thehead of the gannet is controlled and that the serrated beakis facing away from you.

11.4.1 Catching gannets

Gannets are best caught with a gannet crook. This is a light,long (c. 2.5 m) pole, at the end of which is thick but flexiblepiece of wire, bent into a hook shape (Fig. 24). The diameterof the hook should fit comfortably around a gannet’s neck,but must be too narrow for the bird to slide its head out. Makesure there are no sharp edges or points. Approach the gan-net you want to catch, slowly move the hooked end of thepole towards and past the head. This will enable one to pullthe crook onto the neck. Pull the gannet towards you. Whilestill caught in the pole, grab the gannet’s head from behind,before releasing the crook, then use the other hand to holdthe body of the gannet, while keeping its wings tucked in.Alternatively, use the second hand to grab the wings be-tween elbow and shoulder and hold them together behind thegannet’s back (i.e. in the position it assumes when plunge-diving). Be careful not to damage the wings of the bird or thefeet. The webs of gannet feet have an extensive network ofblood vessels close to the skin and therefore tend to bleedeasily.

Figure 24: Diagram of a gannet crook


11.4.2 Breeding population estimates from nest counts andnest densities

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Island map∗∗∗∗∗ Four wooden 2 m poles∗∗∗∗∗ Tape measure or GPS (optional)

At most localities, Cape Gannets nest in large, dense colo-nies, which make accurate ground counts of nests impossi-ble. Vertical aerial photos should be taken of these breed-ing localities at peak breeding, between November andJanuary (Crawford et al. 2007). Peak breeding may differbetween years and localities; this needs to be taken intoaccount when planning an aerial survey. Individual nests areusually easily distinguished on aerial photos, and cannot beconfused with the nests of other seabird species. To ensurethat all colonies at a locality are counted on the photographs,rough outlines of colony position and extent should bemapped from the ground. Individual nests should preferablybe counted on the aerial photos. This is, however, laboriousand time-consuming. If counting individual nests is not fea-sible, it may be more practical to calculate the area occupiedby each colony, by tracing the perimeter of each colony.Ground measurements, for example of straight edges onwalkways or buildings, need to be taken using a tape meas-ure or using a GPS, to provide a scale for the photographs.

Where the breeding population size is estimated from thearea occupied by breeding penguins, colony-specific nestdensities need to be established in the field (i.e. the numberof nests per m2 must be measured) to calculate the numberof nests from area. Within a colony, place four poles, eachmeasuring 2 m, to form a square. Estimate the number ofnests within the square. Nests which only fall partially withinthe quadrat are counted at partial nests; partial nests aresummed to give full nests. Thus, in the example below (Fig.25), the quadrat includes six whole nests and four partialnests (= two full nests), giving a nest density of eight nestsper 4 m2, or two nests per m2 (Crawford et al. 2007). Thismethod is potentially highly disturbing and should only beused at the edge of a colony (where nest densities are, how-ever, likely to be lower, yielding density underestimates) orshortly after the breeding season is finished, when individualnest sites are still recognizable.


Equipment needed:

∗∗∗∗∗ Notebook / nest forms and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Gannet crook∗∗∗∗∗ Balance(s)∗∗∗∗∗ Stopped wing ruler (optional)∗∗∗∗∗ Ruler (optional)∗∗∗∗∗ Vernier, dial or digital callipers (optional)∗∗∗∗∗ Markers for staking out nests (optional)∗∗∗∗∗ Ringing equipment (optional)

The procedure on monitoring breeding success in CapeGannets largely follows the same protocol as for AfricanPenguins (see section 11.1.4). Check and record details ofnest contents of monitored nests from incubation until thechick fledges or the nesting attempt fails. Weekly nestchecks are adequate for general breeding success studies,but should take place every three to five days if chick growthis investigated. To check nest contents, it may be necessaryto gently lift the parent off the nest. Do this by gently push-ing the gannet crook against the sternum of the bird. Gan-net chicks may crèche as they near fledging age. To differ-entiate between study chicks, they may be temporarilymarked with numbered plastic colour rings before replacingthe plastic ring with a permanent stainless steel ring. Ensurethat the colour ring is removed before the bird fledges. Al-ternatively, mark the chick with some dye. Owing to the po-tential disturbance caused by moving through a gannetcolony to handle nest contents, study nests should be cho-sen near the periphery of a colony.

Record nest contents during each visit, with an indicationof chick age. Categorize chick ages similar to that given forAfrican Penguins (see section 11.1.4). Measuring chickgrowth gives an indication of parental feeding effort and foodavailability. Weigh study chicks (under 300 g to the nearest1 g, between 300 g and 1000 g to the nearest 5 g, and above1000 g to the nearest 25 g). Take measurements of studychick culmen length (to the nearest 0.1 mm) and wing length(to the nearest 1 mm). Note the degree of tick infestation oneach chick (i.e. the approximate number of ticks). In order tostandardize the measuring protocol, study chicks should bemeasured in the same order and more or less at the sametime of the day. Chicks are weighed and measured until theyeither die or are completely feathered and ready to fledge.To allow comparison between years, the same sites shouldbe used in subsequent breeding seasons.

11.4.4 Re-sightings of banded individuals

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Gannet crook

Cape Gannets are ringed on the leg, usually with a stainlesssteel ring. Cape Gannets have in the past been ringed withplastic rings in addition to stainless steel rings, particularlyto allow easy visual distinction of cohorts. However, the qual-ity of plastic rings has been found to deteriorate relativelyquickly and eventually break due to material stress broughtabout by plunge-diving. Damaged plastic rings pose an in-jury threat to gannets. The number on the stainless steelband usually cannot be read with the aid of binoculars or atelescope, and a ringed individual needs to be captured.Recapture of ringed individuals may be done opportunis-tically, while carrying out other activities, but regular, sched-uled sessions at staffed localities are advantageous. Scana gannet colony from different angles using binoculars or atelescope. Catch ringed individuals, read the ring numberand record it. Release the bird near where it was caught. Do

Figure 25: Example of how nest density is estimated


not attempt to catch a ringed individual, if it is standing deepin a colony and cannot be hooked from the edge of thecolony or without causing disturbance to surrounding gan-nets.

11.4.5 Diet samples using a stomach flushing technique

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Gannet crook∗∗∗∗∗ Bucket∗∗∗∗∗ Plastic sampling bags (freezer bags)∗∗∗∗∗ Permanent marker pen∗∗∗∗∗ Waterproof labels and pencil∗∗∗∗∗ Ringing equipment (optional)

Cape Gannets tend to regurgitate spontaneously when han-dled, so be sure that the equipment is ready to avoid dietsamples being spilled. Only catch gannets arriving at thecolony. Catch a gannet and immediately invert it over abucket. To reduce potential spillage, pre-place a samplingplastic bag held by a clothes peg in the bucket. Ensure thatanything spilled between the bucket and the sample bag isplaced with the rest of the sample. Hold the bird either be-tween your knees or under one arm. Hold it behind the headwith one hand and support the body with the other hand.Stroke its throat and apply pressure with knees or arms tothe stomach region to induce regurgitation. After the dietsample has been collected, release the individual close towhere it was caught. The individual may be ringed beforerelease. Label the sample bag (species, date, locality, sam-ple number, ring number of bird if applicable) with a markerpen. Add a waterproof label (written on with pencil) with thesame information inside the sample bag, in case the markerpen labelling rubs off while the sample is awaiting process-ing.

Diet samples should be collected weekly or (at least)monthly to identify seasonal changes in diet composition inaddition to inter-annual differences. Always aim to collectsamples at the same time of day. Record all birds caught withempty stomachs and record all cases of fishery related inci-dents of ingested gear, etc.

11.4.6 GPS logger deployment

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Gannet crook∗∗∗∗∗ GPS logger – recharged, started and closed properly∗∗∗∗∗ TESA tape∗∗∗∗∗ Plastic logger outline template∗∗∗∗∗ Scissors∗∗∗∗∗ Scalpel or knife∗∗∗∗∗ Rubber glue (Pattex)

GPS loggers with a weight of 25 g and an accuracy ofc. 10 m are used to track the geographical position and alti-tude of individual gannets. This yields detailed informationon flight path (including that of the out flight, search flight andreturn flight), time spent drifting on the sea surface, the totaldistance covered and the duration of the foraging trip. Themain advantages of this logger are: (a) easy attachment,which minimizes potential disturbance effects, (b) excellentspatial resolution (c. 10 m), (c) adequate sampling frequency(one reading per second), (d) comparatively low cost, (e)possibility of re-use after battery replacement.

Loggers need to be charged before each application and

sealed in waterproof bags. Total logger weight should notexceed 52 g (i.e. less than 2% of the adult body weight).Select a gannet on a nest with an appropriately-aged chickand catch it after its partner returns from a foraging trip. Theage of the chick should be selected depending on the re-search needs. After the greeting ceremony between partnersis complete and the partner is settled on the nest, catch thestudy individual with a gannet crook. Attach the logger to itstail feathers and lower back with Tesa-tape, in a similarmanner as described for African Penguins (section 11.1.7).Release the bird near its nest. The deployment of a loggertakes c. 5 minutes and tagged individuals will usually leavethe colony within minutes. Weigh and measure the chick ofthe tagged adult after a couple of hours, allowing it to firstdigest the food it obtained from the returned parent. Monitorthe nest every 30 minutes starting late afternoon of the sameday. Retrieve the logger as soon as the study bird returnsand download the data onto a computer.

11.4.7 PTT deployment

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Gannet crook∗∗∗∗∗ PTT device – ready to be deployed∗∗∗∗∗ Small cable ties∗∗∗∗∗ Cloth with acetone∗∗∗∗∗ Araldite epoxy (see also section 11.1.8)∗∗∗∗∗ Latex gloves

PTT devices (weighing approximately 20 g) should be usedrather than GPS loggers to obtain information spanning morethan one feeding trip, for example from non-breeding indi-viduals or to document an entire chick-rearing attempt. Po-sition data from PTTs are less accurate than those from GPSloggers (see section 11.1.8 for more information).

Attaching the device requires two people. The personworking with the glue must use latex gloves and should nottouch any part of the bird other than the application area.Test the PTT before application to ensure that it is transmit-ting properly. Remove the magnet to test, using either thetester supplied by the manufacturer.

After capture, secure the bird so that it cannot flap or kickand ensure that the tail is unobstructed. Place a bag over thehead of the bird to keep it calm. Tape paper around the outertail feathers on either side of the two central feathers. Re-move the magnet on the PTT to activate it. Wipe the twocentral feathers near where the feathers are attached to therump with acetone to remove oils that may impair attach-ment. Mix the two components of the Araldite Epoxy (in equalproportions) and add it to the upper surface of the PTT. Placethe underside of the central two tail feathers onto the gluedsurface of the PTT ensuring that the antennae protrudesupright from between them. Add a little more glue to theupper surface of the feathers to improve the attachment. Fita small cable tie around each end of the PTT for security ofattachment. Pull cable ties tight, taking care not to split thefeathers in doing so. A ring can be fitted before release and/or a dye painted onto visible parts of the bird. When the glueis dry, trim the cable ties, remove the bag and release thebird. Allow the bird to vomit if it needs to while applying thePTT.

11.5 White-breasted Cormorant (Phalacrocorax lucidus)

Adult White-breasted Cormorants are easily distinguishedfrom other cormorant species by their large size and white


throat and breast. The entire underparts are white in imma-ture birds. Juvenile individuals resemble immature individu-als but throat and upper neck is brown. White-breasted Cor-morants are gregarious at roosting and breeding sites, andoften associate with other cormorants or pelicans. They gen-erally nest colonially. Breeding White-breasted Cormorantsare highly susceptible to human disturbance.

11.5.1 Breeding population estimates from nest count

Equipment needed:


areas∗∗∗∗∗ Tally counter

Counts of active nests (i.e. nests containing eggs or chicks)should be undertaken monthly, particularly at localities whereWhite-breasted Cormorants breed year-round. If possible,separately record the number of birds incubating or brood-ing and the number of nests with chicks (and how manychicks). At localities where this is not feasible, a number ofcounts spanning at least a month either side of the knownbreeding peak at that locality are necessary to provide anaccurate estimate of the breeding population. Additionalcounts of active nest sites must be recorded separately andclearly indicated. Active nest sites do not contain eggs orchicks, but include nests being built or having recently beenconstructed, sites defended by at least one adult and sitessupporting courting pairs.

Always count colonies from a distance to prevent nestabandonment. Nests containing eggs or small chicks are rec-ognized by the adult sitting deep in the nest.


Equipment needed:


areas∗∗∗∗∗ Colony map or photo indicating study nest

∗∗∗∗∗ Catching hoop-net (optional)∗∗∗∗∗ Ringing equipment (optional)

Owing to the sensitivity of White-breasted Cormorant colo-nies to disturbance, monitoring of study nests to determinebreeding success should only be done from a distance.Choose a number of study nests while the pair is still nestbuilding and courting to allow an estimation of the start ofincubation. To make sure that the same nest is checked eachweek, draw a colony map or photograph the colony, and in-dicate the study nests. Observe nest contents from a dis-tance – to see the nest contents, wait for the adult to shift onthe nest or for the parents to change over. Do not disturb thenest by physically handling adults or nest contents, as thismay lead to nest abandonment. If nest contents are lost, tryto establish the cause of mortality.

Fledglings may be caught with a light hoop-net (Fig. 26)and ringed. Avoid disturbance to other breeders. If possiblefit colour rings in addition to steel rings. Colours should beallocated to identify individual cohorts and breeding localities(see section 4).

11.6 Cape Cormorant (Phalacrocorax capensis)

Cape Cormorants are gregarious, roosting and breeding inlarge groups. They are distinguished from Bank Cormorantsby their smaller size and more slender appearance and bythe yellow facial skin and gular pouch. Juvenile and imma-ture individuals are browner and paler. After leaving the nest,fledglings remain with their parents for several weeks. CapeCormorants are particularly susceptible to human distur-bance and readily abandon nests. In extreme cases, onepanicked single bird may cause mass abandonment. Specialcare must be taken to avoid any disturbance; this makesmonitoring efforts difficult.

11.6.1 Breeding population estimates from nest counts anddensities

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Locality map∗∗∗∗∗ Four wooden 2 m poles∗∗∗∗∗ Tape measure or GPS (optional)

Depending on the size of the colony, different strategies toestimate the breeding population may be used. Groundcounts, following the method outlined for White-breastedCormorants (section 11.5.1), are suitable for small colonies(<1000 nests). Several counts around the putative breedingpeak should be undertaken.

For larger colonies, single annual counts using aerial pho-tography may be the only way to obtain accurate estimates(Shelton et al. 1982). Counts should be done as close to thepeak of the breeding season as possible. This peak may varybetween localities and years. Methods detailing aerial pho-tography, ground measurements for scaling purposes andnest density measurements should follow those described forCape Gannets (section 11.4.2). In addition to estimating nestdensities, the extent of colonies, particularly in the case ofseveral cormorant species nesting together, needs to becarefully mapped. At localities where ground counts are pos-sible, count and record all unattended chicks congregated ingroups. Divide the total by three to estimate the number ofnests (Shelton et al. 1982).

Figure 26: Example of a hoop net used to catch cormorants, gullsand terns



Cape Cormorant colonies are very easily disturbed and anymonitoring of study nests to determine breeding successshould only be done from a distance, following the methodgiven for White-breasted Cormorants (section 11.5.2).

11.6.3 Diet samples (from pellets)

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Drying tray∗∗∗∗∗ Tissue paper∗∗∗∗∗ Masking tape∗∗∗∗∗ Sampling bags∗∗∗∗∗ Marking pen

Cape Cormorants regurgitate pellets made of food wastesuch as fish bones. Otoliths contained in these pellets givean indication of diet composition. Collect fresh pellets atknown roost sites. Where Cape Cormorants nest togetherwith Bank Cormorants, pellets can be distinguished by theirsmaller size. Fresh pellets may be surrounded by a layer ofyellowish mucous in Cape Cormorants; those of Bank Cor-morants are covered in clear mucous. Pellets of Cape Cor-morants tend to be smaller than those of other cormorants.

Collect pellets individually, noting date and place of col-lection. Assign a number to each pellet. Place pellets on adrying tray. This may be a simple tea tray or one with sepa-rate compartments (like a printer’s tray) to help keep track ofindividual pellets. Do not mix up pellets. Dry pellets in thesun. Pellets may also be dried in a drying oven at low tem-peratures. This is, however, not recommended as it maydamage otoliths. Make sure that pellets are completely drybefore packing them for further analysis because they mightget mouldy. If pellets cannot be processed straightaway, rollup individual pellets in tissue paper and tape tissue paperwith masking tape to prevent it from unravelling. Mark thepellet number on the masking tape. Pack sets of samples(per date and locality) in separate sampling bags, in whicha label with all the information pertaining to numbered pel-lets is contained. Store sample bags in a dry place until pel-lets can be analysed. If pellets can be pre-processedstraightaway, line up the extracted otoliths. Drop a sectionof sticky tape onto the row of otoliths and fold the tape overto seal the sample. The strips of sticky tapes containing theotoliths must be labelled with a permanent marker pen be-fore storing the samples for later analysis. Collect pelletsthroughout the year to allow an analysis of diet seasonality.

11.6.4 Diet samples (from regurgitation)

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Plastic sampling bags (freezer bags)∗∗∗∗∗ Waterproof labels and pencil∗∗∗∗∗ Permanent marker pen

Cape Cormorants may regurgitate on landing, if approachedtoo closely or if handled. Because regurgitation is a stressresponse, samples should only be collected opportunistically(e.g. if fledglings regurgitate spontaneously while beingringed) and not as part of a standard routine. Collect thesample in sample bags, label the outside of bags with amarker pen (species, date, locality, age class of individual)and freeze bag until the sample can be processed. Add awaterproof label with the same information written in pencil

inside the bag. Take note that not necessarily the entirestomach contents are regurgitated; data obtained fromregurgitations therefore yield information on the identity ofitems found in the diet, but not necessarily on the percent-age contribution of these items to the diet.

11.7 Bank Cormorant (Phalacrocorax neglectus)

Bank Cormorants are heavily built, glossy black, with a whiterump when breeding. Juvenile and immature individuals area dull black., with dark brown or blue-grey eyes. Adults havetwo-toned eyes, with an orange to brown top half and a greenbottom half Bank Cormorants nest in colonies of c. 20–100pairs, sometimes adjacent to colonies of other seabirds andoften roost with other cormorants. After leaving the nest,fledglings remain with their parents for up to three months.

11.7.1 Breeding population estimates from nest counts

Counts of active nests and of active nest sites should follow-ing the methods outlined for White-breasted Cormorants(section 11.5.1).


Bank Cormorant colonies are very easily disturbed and anymonitoring of study nests to determine breeding successshould only be done from a distance, following the methodgiven for White-breasted Cormorants (section 11.5.2).


The procedure for collecting, labelling, drying and storingBank Cormorant pellets follows that outlined for Cape Cor-morants (section 11.6.3). Note that Bank Cormorant pelletsare larger than those of Cape Cormorants and are sur-rounded by a layer of clear mucous when fresh.

11.8 Crowned Cormorant (Phalacrocorax coronatus)

This small cormorant is recognized by its small size, red eyesand partially erect frontal crest. Juvenile individuals are darkbrown rather than glossy black and lack a crest and haveyellow eyes. Crowned Cormorants breed in small groups,often in association with other seabird breeding colonies.Colony location may vary between years and breeding maytake place throughout the year. Breeding is relatively wellsynchronized at the colony, but not necessarily at the local-ity scale. Crowned Cormorants readily abandon nests if dis-turbed.


Counts of active nests and of active nest sites should follow-ing the methods outlined for White-breasted Cormorants(section 11.5.1).


Crowned Cormorant colonies are very easily disturbed andany monitoring of study nests to determine breeding successshould only be done from a distance, following the methodgiven for White-breasted Cormorants (section 11.5.2).

11.8.3 Diet samples

Collect regurgitations opportunistically and process them in


the same manner as described for other cormorants (seesection 11.6.4).

11.9 African Black Oystercatcher (Haematopus moquini)

African Black Oystercatchers are easily recognizable by theirblack plumage, long red legs and beak, and a prominentfleshy red eye ring. Adults are mostly sedentary and territo-rial throughout the year. They are solitary nesters. Nests areoften within 3 m of the high-water mark. Oystercatchers areeasily disturbed during the breeding season. Small chickshide in response to parents’ alarm call and can be difficult tofind.

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod (optional)∗∗∗∗∗ Vernier, dial or digital callipers (optional)∗∗∗∗∗ Portable electronic balance (optional)∗∗∗∗∗ GPS∗∗∗∗∗ Soft (“B”) pencil

11.9.1 Year-round monitoring

At regular intervals, say every spring tide or alternative springtides, undertake counts of oystercatchers on the entire is-land, demarcated study area or fixed section of mainlandshoreline. Note whether birds are in pairs, single, or a group.If feasible, take the GPS positions near the high-tide level atright angles to the position of each oystercatcher (or pair/group of oystercatchers) on the shore. This detailed mode ofdata collection provides a methodological opportunity todevise a fairly sophisticated way to analyze spatial countdata (e.g. not just documenting the increase but looking atthe process of where any additional birds were accommo-dated).

11.9.2 Breeding success

Oystercatchers lay their eggs in inconspicuous scrapes;nests may therefore be difficult to locate (Fig. 27). Startmonitoring a few weeks prior to the normal start of the breed-ing season at the locality. Find nests as soon as feasible afterlaying, by observing breeding pairs from a distance and not-

ing the general position of the nest. Once pairs are disturbed,they will move away from their nest, making nests more dif-ficult to find. Approach the area cautiously, taking care notto accidentally step onto the nest and record the GPS coor-dinates for the nest; these can be used to relate nest posi-tions to those of previous years (and second and third at-tempts at relaying within a season).

The intensity of breeding success monitoring activities willdepend on the exact needs of research questions posed, butcare should be taken to keep invasive activities to a mini-mum. Basic breeding success parameters such as egg-lay-ing date, incubation length, number of eggs, number of fledg-lings, fledging period and time to final departure may beobtained from watching the nest and observing parentalbehaviour from a distance using a telescope.

Methods for more intensive monitoring:

Weigh and measure the eggs (see section 3.1). Mark theeggs, using a soft pencil to recognize individual eggswhen re-weighing them.

Monitor nests at 3–5 day intervals to check for egg loss.If possible do this from a distance, using binoculars ora telescope. For nests which failed, try to determinecause of egg loss. If possible, reweigh eggs after 14days, and after 21 days. These measurements can beused to confirm rates of mass loss during incubation.

Determine how many eggs hatch. Monitor chicks at 3 to5 day intervals. Parents will be particularly vocal and agi-tated after eggs hatch, and small chicks can be ex-tremely difficult to locate. If chicks can be located, takestandard measurements (culmen, total head, tarsus,foot, wing, tail). These measurements can be used toassess chick condition. Ring chicks with stainless steelbands the first time they are encountered after theyreach a weight of 140 g. Try to determine ages at whichchick losses occur (measure and weigh fresh carcass).Note if and when the pair relays after losing an egg orchick. Determine the fledging period and the period be-tween fledging and final departure (about 100 days).

11.10 Kelp Gull (Larus dominicanus vetula)

Adult Kelp Gulls are white with black wings; juveniles arespeckled brown to grey, becoming paler as they get older.Kelp Gulls may nest solitarily or in loose to moderately densecolonies. They aggressively defend their territories and nestsfrom intruders (including humans), diving and striking at in-truders with their beak and feet while emitting a high-pitchedscreech, and may defecate or regurgitate on an intruder.Wear protective clothing, including a hat when moving nearor between nests. The disturbance caused by upsetting abreeding Kelp Gulls may trigger further disturbance of othernesting Kelp Gulls or of colonies of other seabirds breedingnearby. This may lead to mass nest abandonment in extremecircumstances. Kelp Gulls predate exposed nest contents(eggs or small chicks) of disturbed breeders, including thoseof other Kelp Gulls. It is therefore vital to keep disturbanceof breeding Kelp Gulls to a minimum if there is a possibilityof disturbing other birds.


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)

Figure 27: A cryptic African Black Oystercatcher nest (arrow) onHalifax Island, Namibia


∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod∗∗∗∗∗ Tally counter

Serial counts spanning at a month either side of the sus-pected breeding peak at that locality are necessary to pro-vide an accurate estimate of the breeding population. KelpGull chicks are semi-precocial and can move from the nestwithin hours after hatching. Egg-laying is relatively well syn-chronized and counts should therefore be done during thetime when most individuals are incubating. Record nestsbeing incubated or brooded separately from free-runningchick. Count each chicks as a nest (Crawford et al. 1997).Always count colonies from a distance to prevent nest aban-donment and egg predation.


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Tally counter∗∗∗∗∗ Insulation tape and scissors (optional)∗∗∗∗∗ Bird marking dye (optional)∗∗∗∗∗ Ringing equipment (optional)∗∗∗∗∗ Balance (optional)∗∗∗∗∗ Ruler and callipers (optional)

For general breeding success estimation (i.e. the number ofchicks fledged per pair) it is sufficient to identify a study areawith 50 to 100 nests. Check how many eggs are laid in totalin the study area. Check again how many chicks hatchedand how many of the chicks fledge successfully.

Because Kelp Gull chicks are highly mobile and breedersare susceptible to disturbance, following individual studynests in detail (for example to establish chick survival orgrowth rates) is not easy. Mark both parents with colour ringsand possibly with a spot of dye while they are incubating.This will make it easier to find the chick again. Mark thehatchling(s) on the leg with a temporary band made from ashort section of coloured insulation tape. Cover hatchlingswith nesting material to prevent them from being predated byother gulls. Watch from a distance to ensure that the parentresettles on the nest. Monitor nest at regular intervals, atleast weekly, but preferably every five days. Make sure thatthe insulation tape is not becoming too tight as the chickgrows. Replace the tape if necessary. Catch the chick witha hoop net (Fig. 26) before it starts flying (roughly when it hasshed a quarter of its downy feathers). Remove the tape andfit a stainless steel ring.

11.10.3 Ringing and recapture

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Suitable walk-in trap∗∗∗∗∗ Ringing equipment∗∗∗∗∗ Artificial eggs∗∗∗∗∗ Field incubator or cooler box with towel

Adult Kelp Gulls are best caught for ringing or recapture witha walk-in trap placed over a nest containing eggs. Ring bothmembers of a pair, preferably with a colour ring in additionto the stainless steel ring, to investigate factors such as matefidelity, nest site fidelity, and incidence of replacement lay-ing. To ensure that eggs are not damaged while extractingthe parent from the trap, replace the clutch with artificial eggs.

Keep the clutch safe and warm in a field incubator (orwrapped in a towel inside a cooler box). Replace the clutchin the nest as soon as the parent has been ringed and re-leased.


The procedure for collecting, labelling, drying and storingKelp Gull pellets follows that outlined for Cape Cormorants(section 11.6.3). Note that Kelp Gull pellets are more looselypacked than those of Cape or Bank Cormorants. They tendto disintegrate relatively quickly, making them difficult to rec-ognize if they are not fresh. In addition to providing informa-tion on seasonal differences in diet, Kelp Gull diet samplesgive an indication of the percentage of the diet consisting offish factory offall or rubbish.

11.10.5 Diet samples (from regurgitations)

Kelp Gulls may spontaneously regurgitate when handled.Methods for collecting regurgitation samples follow thosedescribed for Cape Cormorants (section 11.6.4).

11.11 Hartlaub’s Gull (Larus hartlaubii)

Adult Hartlaub’s Gulls are distinguished by a pale grey backand upper wings, white underparts and dark red legs andbeak. In breeding adults, the crown, upper nape and throatare faded pale grey, often only visible as a faint dark outline.The head of non-breeding adults is white. The head andupper parts of juvenile (first year) birds are mottled lightbrown. Hartlaub’s Gulls breed colonially, often in mixed colo-nies with Swift Terns and sometimes with Grey-headed Gullsor Crowned Cormorants. They may be displaced from theirnests by breeding Swift Terns. Breeding tends to be oppor-tunistic. Breeding seasonality and colony location are poorlydefined and may differ between localities and years. Egg-laying tends to be poorly synchronized within colonies.

Hartlaub’s Gulls are aggressive and vocal when breeding,but rarely physically attack humans as Kelp Gulls do.Hartlaub’s Gulls predate exposed eggs of cormorants, ternsand other Hartlaub’s Gulls. Keep disturbance of breedingHartlaub’s Gulls to a minimum if there is a possibility of dis-turbing other birds, particularly if they are breeding in mixedcolonies.


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Telescope and tripod∗∗∗∗∗ Tally counter

Monthly counts, or if not possible, several counts spanningat least a month either side of the suspected breeding peakat each locality are necessary to provide an accurate esti-mate of the number of breeding Hartlaub’s Gulls. Countsshould be made during the time when most individuals areincubating. Separately record the number of nests being in-cubated, brooded or containing chicks, as well as free-running or crèching chicks. Divide the number of crèchingchicks by two to estimate the minimum number of breedingpairs they represent (Williams et al. 1990). Because nestdensity is generally high, nest counts are best done from anelevated vantage point. Count colonies from a distance toprevent nest abandonment and egg predation. Take care to


detect Hartlaub’s Gull nests in mixed colonies with SwiftTerns.


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Colony map∗∗∗∗∗ Nest markers (optional)∗∗∗∗∗ Ringing equipment (optional)∗∗∗∗∗ Hoop net (optional)∗∗∗∗∗ Herding screens (optional)∗∗∗∗∗ Temporary fencing (optional)

Breeding Hartlaub’s Gulls (and Swift Terns or Crowned Cor-morants if nesting in mixed colonies) are prone to distur-bance. Only general breeding success should be estimated.Because egg-laying may not be well synchronized, map astudy area. Visit the study area at least weekly. From a dis-tance, count and separately record the number of nests be-ing incubated or brooded, free-running chicks and thenumber of fledglings in the study area.

At regularly used breeding localities, mark a set of nestsites prior to breeding. Choose nest sites that are easily vis-ible from a distance.

Unless trapping and ringing activities are likely to causedisturbance to other breeders, including to other species,Hartlaub’s Gull chicks may be caught before becomingflighted using a hoop net (see Fig. 26) or by herding groupsinto a temporary enclosure.


The procedure for collecting, labelling, drying and storingHartlaub’s Gull pellets follows that outlined for Cape Cormo-rants (section 11.6.3). Pellets from Hartlaub’s Gulls are largerthan those from Swift Terns, although sizes may overlap.

11.11.4 Diet samples (from regurgitations)

Hartlaub’s Gulls may spontaneously regurgitate when han-dled. Methods for dealing with regurgitation samples followthose described for Cape Cormorants (section 11.6.4).

11.12 Swift Tern (Sterna bergii)

A large tern with characteristic black cap and crest and yel-low beak. Non-breeding adults have a white forehead and aspotted or streaked crown. Juveniles have a mottled darkmantle, back and upper wings. Non-breeding individuals maymove substantial distances. Swift Terns are usually colonialbreeders, but may nest in sparse numbers in breeding colo-nies of Hartlaub’s Gulls, Crowned Cormorants, Grey-headedGulls and Roseate Terns. Nest density is high. Egg-laying issynchronous within colonies. Chicks are semi-precociousand remain at the nest for about four days. Older chicks mayform crèches. Breeding Swift Terns are less aggressive thanHartlaub’s Gulls and nest contents, particularly chicks, arevulnerable to predation by Kelp Gulls and Great White Peli-cans.


Counts of nests should following the methods outlined forHartlaub’s Gulls (section 11.11.1). In addition to occupied

nests, count each free-running or crèching chick. Each chickrepresents a breeding pair (Cooper et al. 1990).


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ Colony map∗∗∗∗∗ Nest markers (optional)∗∗∗∗∗ Ringing equipment (optional)∗∗∗∗∗ Herding screens (optional)∗∗∗∗∗ Temporary fencing (optional)∗∗∗∗∗ Cardboard boxes (optional)

Measure breeding success following the methods describedfor Hartlaub’s Gulls (section 11.11.2). Breeding in Swift Ternstends to be well synchronized at colony level and chickstherefore tend to fledge more or less at the same time. Oncechicks start forming crèches and before they are able to fly,herd and surround groups with herding screens into a tem-porary holding enclosure made from temporary fencing.Transfer the chicks into cardboard boxes, where they aremore likely to remain calm. Ring, weigh and measure allcaptured chicks before transferring the birds from the boxesback into the enclosure and remove the temporary fencing(Le Roux 2002).


The procedure for collecting, labelling, drying and storingSwift Tern pellets follows that outlined for Cape Cormorants(section 11.6.3). Pellets from Swift Terns are smaller thanthose from Hartlaub’s Gulls; sizes may overlap.

11.12.4 Diet samples (from direct observation)

Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars

Swift Terns carry whole prey items back to their chicks intheir beak. Position yourself as close as possible to thebreeding colony without disturbing it. Scan terns returning tothe colony carrying fish in their beaks. Using binoculars,attempt to identify the prey and estimate the length of eachitem by comparing it to the length of the bird’s culmen(Crawford and Dyer 1995). This method only allows the iden-tification of prey items and their approximate sizes but doesnot allow the quantification of the percentage contribution ofthese to the diet.

11.13 Damara Tern (Sterna balaenarum)

A small tern with a characteristic call. Breeding adults havea black cap extending well below the eye; in non-breedingadults the forehead to mid-crown is white and mottled palegrey. Juveniles have brown barring on upper parts. DamaraTerns nests solitarily or in loose groups (inter-nest distance>100 m) on flat gravel plains, salt pans or on flat sand in dunefields. Breeding seasonality varies between localities. Breed-ers (singly or in groups) mob intruders (including humans)approaching nest or free-running chick, but do not attack ifnest contents are handled.



Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ GPS

The cryptic nature of Damara Tern nests and the habit ofbreeding adults to fly off the nest to mob an intruder, even ifthe intruder is still hundreds of metres away from the nest,make it difficult to conduct nest counts. If nest sites positionsfrom previous years are known, check the general vicinity forsigns of activity. At localities where nest sites are suspected,walk around the general area, listening for the tern’s distinc-tive call and taking care not to step on a nest. When you heara Damara Tern call, sit down and locate the calling tern withbinoculars. Remain sitting and follow the Damara Tern withbinoculars until it lands and settles. Using any nearby land-marks as reference points, pinpoint the location of the birdand walk to the site. As you get close to the site, carefullyscan the ground for a shallow, inconspicuous scrape contain-ing one egg or a semi-precocious chick which may hide orfreezes if caught in the open. Take care not to inadvertentlystep on an egg or chick. Take a GPS position of the egg orchick (see Fig. 28). If a suspected Damara Tern nest is dis-covered by chance during a nest search, take the nest’s GPSposition and retreat. With binoculars, confirm the identity ofthe nest by watching parents returning to the nest. DamaraTern nests may easily be confused with those of other spe-cies, particularly White-fronted Plovers or Chestnut-bandedPlovers. Damara Terns lay one egg which is generally moreelongate than that of plovers.


Equipment needed:

∗∗∗∗∗ Notebook and pen(cil)∗∗∗∗∗ Binoculars∗∗∗∗∗ GPS∗∗∗∗∗ Spring balance and cloth bag or portable electronic bal-

ance∗∗∗∗∗ Vernier, dial or digital callipers∗∗∗∗∗ Ringing equipment (optional)

Once a nest is located and the position has been recorded,carefully weigh and measure the egg (see section 3.1). Ide-ally, visit the nest every third day to check for egg loss andreweigh the egg each time. These measurements can beused to confirm rates of mass loss during incubation. Fornests which failed, try to determine cause of egg loss.

After the chick hatches, monitor it at three to five day in-tervals. If chicks can be located, take standard measure-ments each time it is found to assess chick condition. Ringthe chick with a stainless steel ring while still small (afterabout a week), before it becomes too mobile. Try to deter-mine ages at which chick losses occur (measure and weighfresh carcasses). Record when the chick is able to fly andwhen it finally leaves the area. Note if and when the pairrelays after losing an egg or chick.

Diet samples (from direct observation or regurgitation)

Damara Terns take whole food items back to their chicks.Position yourself near the nest site, but not too close to dis-turb the nest. With binoculars attempt to identify the prey itemgauge its length using the culmen of the bird as an indica-tion.

Chicks may spontaneously regurgitate when being han-dled. Methods for collecting regurgitation samples followthose described for Cape Cormorants (section 11.6.4). Dietdata obtained from either method only yields information onthe identity of prey items, but not on the percentage contri-bution of these items to the diet.

Acknowledgements – This paper is a contribution to the project LMR/EAF/03/02 of the Benguela Current Large Marine Ecosystem(BCLME) Programme. Thank you to K. Peard for scanning the draw-ings and some of the photographs. All drawings and photographsare by the author with the exceptions of Figs 16–23 (R. Jones) andFig. 3 and part of the cover collage (R. Mullers).

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Annex 3 MONITORING SEABIRDS IN THE BCLME: DATA …