© 2002 by The Society for Integrative and Comparative Biology
| ||||||||||||||||||||||||||||||||||||||||||||||||||||
From Odor Molecules to Plume Tracking: An Interdisciplinary, Multilevel Approach to Olfaction in Stomatopods1
1 Department of Integrative Biology, VLSB 3060, University of California, Berkeley, Berkeley, California 94720-3140
 |
SYNOPSIS |
|---|
TOP
 SYNOPSIS INTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
Like many marine crustaceans, mantis shrimp rely on their sense of smell to find food, mates, and habitat. In order for olfaction to function, odorant molecules in the surrounding fluid must gain access to the animal's chemosensors. Thus fluid motion is important for olfaction, both in terms of the large scale fluid movements (currents, waves, etc.) that advect the odorants to the vicinity of the sensors, and the small-scale viscosity dominated flows that determine odorant access to the surface of the sensor. In order to understand how stomatopods interpret their chemical environment, I investigated how stomatopod chemosensory morphology and the movement of the structures bearing the chemosensors affect fluid access to the sensor surface in Gonodactylaceus mutatus. Preliminary results from new directions are presented, including mathematical modeling of molecular flux at the sensor surface, field studies of the effects of ambient flow on odor sampling behavior, and flume experiments testing the ability of stomatopods to trace odor plumes. Finally, I show how the use of multiple techniques from several disciplines leads to new ideas about the functional morphology of stomatopod antennules.
|
INTRODUCTION |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
Many marine organisms are subject to complicated chemical signals that vary in time and space at multiple scales. Examining how stomatopods interpret their chemical environment, sort out the relevant olfactory cues, and navigate up odor plumes will give us insight into the general problems of odor signal recognition and search strategy.
Stomatopods
Stomatopods are shrimp-like crustaceans (Fig. 1A) that live in burrows in mudflats or in coral reef rubble in tropical and semi-tropical habitats. Stomatopods are excellent subjects of chemosensory studies because they use olfaction to detect prey items, to find mates and to recognize conspecifics (Caldwell, 1979
, 1985, 1987; Caldwell et al., 1989). Their simple chemosensory appendages facilitate physical and mathematical modeling, and they are available in a large range of body sizes. This study focuses on the Hawaiian stomatopod Gonodactylaceus mutatus with some data from the Californian stomatopod Hemisquilla ensiguera californica.
|
Odor molecules and fluid flow
Odors in the marine environment consist of patches or filaments of high concentrations of odorant molecules (often small molecules, such as amino acids) in the surrounding fluid (Weissburg, 2000
; Crimaldi and Koseff, 2001). In order for olfaction to function, odorant molecules must gain access to the animal's chemosensors. Sensor morphology, arrangement, and movement relative to the ambient water motion affect the flow of water around the sensilla (Gleeson et al., 1993, Koehl, 1995, 1996; Mead and Koehl, 2000). Environmental flows such as waves (Mead, unpublished data) and currents (Weissburg and Zimmer-Faust, 1993, 1994; Moore et al., 1994) affect the relative velocity of the sensor and the fluid and thus are likely to affect the flow of water around the sensilla. This in turn affects the transport of odorants to the chemosensor surface and to the chemoreceptors within the sensilla (Stacey et al., 2002). |
STOMATOPOD CHEMOSENSOR MORPHOLOGY |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
External morphology and arrangement
Stomatopod antennule ablation experiments (Caldwell, unpublished data) suggest that mantis shrimp detect odors from distant sources (such as food or conspecifics) by means of specialized chemosensory sensilla called aesthetascs (Laverack, 1988
; Hallberg et al., 1992; Mead and Weatherby, 2002). The aesthetascs are arranged on the distal dorsal surface of a robust filament that emerges from the base of the lateral filament of the antennule (Fig. 1B). G. mutatus aesthetascs are long, slender cuticularized structures that are enlarged proximally and are inserted into the antennule filament at an angle of about 50 degrees, in rows of three, with one row per filament segment (Fig. 1C). This simple arrangement is in marked contrast to the complex arrays of aesthetascs and associated setae found in most other crustaceans (Derby, 1982; Derby et al., 1997; Gleeson, 1982; Gleeson et al., 1993; Grünert and Ache, 1988; Hallberg et al., 1992). Also unusual is the scaling of the aesthetasc array with body size: as G. mutatus grow in size from 8 mm to 52 mm rostrum-telson length, the number of aesthetasc rows increases from 3 to 18, the aesthetascs increase in length from 200 to up to 550 µm in length and increase in diameter from 10 to 20 µm (Mead et al., 1999).
Internal morphology
Dye studies show that the aesthetasc cuticle is readily permeable to methylene blue (and thus probably small odor molecules as well) along the entire length of the aesthetasc (Mead et al., 1999). Transmission electron micrographs indicate that each aesthetasc is innervated by bipolar sensory cells (Mead and Weatherby, 2002). Basally the aesthetasc cuticle is thick (up to 2 µm) and lamellar. In the distal portion of the aesthetasc, the sensory cells' outer dendritic segments (where the presumptive chemoreceptors are located) branch extensively until there is only one microtubule per membrane sheath (Fig. 1D). Up to 2,500 of these fully branched outer dendritic segments extend to near the tip of the aesthetasc and are the primary contents of the outer 80% of the aesthetasc. Here the cuticle is 0.5 µm thick, has lost its lamellar appearance, and is less electron-dense.
Odor molecules that make contact with the distal portion of the aesthetasc are more likely to be detected efficiently than odor molecules that come into contact with the base (Mead and Weatherby, 2002). The fourfold thinner cuticle at the tip than at the base is likely to result in a 16 times shorter diffusion time (Berg, 1983). In addition, since the amount of surface area available to receptors is much greater in the outer portion of the aesthetasc, diffusion times within the sensilla lumen are likely to be shorter at the aesthetasc tip than at the base.
|
FLUID ACCESS TO SENSORS |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
Odor sampling by antennule flicking
Like many crabs, lobsters, and other crustaceans (Snow, 1973
; Schmitt and Ache, 1979; Atema, 1985; Moore et al., 1991; Gleeson et al., 1993; Koehl, 1995, 1996; Gleeson et al., 1996; Hallberg et al., 1997), stomatopods flick their antennules, probably to increase the velocity of their antennules relative to the surrounding fluid and reduce the boundary layer thickness (Mead et al., 1999). This action increases the volume of odor-containing fluid that can penetrate the aesthetasc array, and brings the fluid closer to the aesthetasc surfaces, minimizing the distance over which the odor molecules have to diffuse before encountering the aesthetasc. This decreases the time required for odorant molecules to diffuse to the sensillar surface, and improves the speed and accuracy with which changes in odorant concentration are reflected at the sensillar surface (Koehl, 1996; Mead and Koehl, 2000). Although it is not known what odor signal characteristics are important to stomatopods, by analogy to lobsters and insects (Kaissling 1998a, b; Gomez et al., 1999; Rospars et al., 2000) it is probable that chemical concentration and chemical flux are signal features likely to be important to stomatopod olfactory receptors.
Reynolds number
One method of investigating how fluid moves around olfactory appendages during flicking is to calculate the Reynolds number (Re) describing the flow around the aesthetascs. Re is a nondimensional parameter that measures the relative importance of inertial and viscous forces for a given flow and is thus a major determinant of flow pattern (Vogel, 1994).
|
|
where 
is the density of the fluid, here the density of sea water at 25°C, 1,023 kg/m3; L is a spatial scale, here the aesthetasc diameter; U is the antennule velocity, and µ is the dynamic viscosity of the fluid; here the viscosity of sea water at 25°C, 0.97 x 10–3 Pa sec. For a given aesthetasc diameter, a large Re indicates that the boundary layer is thin relative to the aesthetasc dimensions (inertial forces dominate), and a small Re suggests that the boundary layer is thick relative to the body (viscous forces dominate).
Antennule velocities were determined by filming G. mutatus olfactory flicks with high speed video while the animals were stimulated with brine shrimp odor (Mead et al., 1999). We found that as animals grow in size from 8 mm to 52 mm rostrum-telson length, the tangential velocity of the antennule tip during a flick outstroke increases from about 2.5 cm/sec to 10 cm/sec, and the tangential velocity of the antennule tip during a flick return stroke increases from about 1.3 cm/sec to about 5 cm/sec. Since aesthetasc diameter also increases with body size, the Re describing the flick outstroke increases from 0.2 to 1.8, and the Re describing the flick return stroke increases from 0.1 to 0.9 over the same range of body sizes. These Reynolds numbers are in a range where (for this geometry) a small change in Re can result in a large change in the volume of fluid that is able to penetrate the aesthetasc array (Cheer and Koehl, 1987; Hansen and Tiselius, 1992; Koehl, 1995). These results suggest that more fluid flows through the aesthetascs during the flick outstroke than on the return, and that more fluid is sampled per aesthetasc by large animals during flicking than by small animals.
Physical model experiments
To test these hypotheses, I constructed physical models of several segments of the antennule filament (Fig. 1E) and its associated aesthetasc rows from juvenile and adult G. mutatus and towed them through a tank of viscous fluid (Mead and Koehl, 2000). Although there is a discrepancy between the linear motion of the model and the rotational movement of the real antennule, the rotational angle of the antennule as it moves through the flick is small (less than 20°; Mead, unpublished data). Linear motion is a reasonable first approximation of antennular motion. By choosing model spatial scales, model towing velocities, and tank fluid viscosities such that Re is conserved, it is possible to analyze flow patterns around a model knowing that the same flow patterns are present when the real animal flicks its antennules in its native habitat. One advantage of this approach is that it is possible to choose model lengths and velocities that facilitate data collection, adjusting the fluid viscosity accordingly. In addition, the use of models makes it possible to test parameter combinations not found in nature.
The models were dragged through a viscous liquid seeded with neutrally buoyant reflective particles. A horizontal light sheet illuminated the particles as they moved around the array of chemosensory sensilla. Flow patterns around the model were recorded and used to generate velocity profiles around the aesthetascs. Flow fields at different distances from the filament were generated by moving the model up or down relative to the light sheet. The ability of fluid to flow through adjacent rows of aesthetascs (leakiness) and the volume flow rate between rows of aesthetascs were calculated as described in Mead and Koehl (2000).
Fluid access to the aesthetasc surface during a flick
Figure 2 shows velocity profiles through the aesthetasc array from the physical model experiments, scaled to the animal. In both the juvenile and the adult, the mean velocity of the fluid moving through the aesthetasc array is several times greater during the outstroke than during the return stroke. Fluid moves three times faster through the adult array than through the juvenile array. Figure 3 shows how fluid velocity varies with distance along the aesthetasc. The mean fluid velocity moving through the array is greater near the tip of the aesthetascs than near the aesthetasc bases because of the boundary layer created by the supporting antennule filament. Thus fluid (and odor molecule) access to the aesthetasc is fastest and greatest in volume along the outer portion of the aesthetasc where the cuticle is thinnest and the branching of the outer dendritic segments is most evolved.
|
minimum significant differences (MSDs). The MSD is a composite of the standard error and the critical value of the studentized range Qcrit (Sokal and Rohlf, 1995
|
Velocity profiles can also be used to calculate volume flow rate. As G. mutatus grow, changes in antennule velocity, aesthetasc length, and number of aesthetasc rows result in 20 times as much fluid per unit time flowing between adjacent rows of sensilla in adults as in juveniles (Mead and Koehl, 2000
). Figure 4 shows the volume flow rate ratio as a function of the Re of the outstroke. Both adults and juveniles sample 5–7 times more fluid per second between adjacent aesthetasc rows during the outstroke than during the return stroke. Changing the Re at which flicking occurs in either juveniles or adults would decrease the difference in flow rates between the two strokes.
|
Stroke asymmetry
The difference in volume flow rates between the two strokes of the flick is likely to have functional importance since stomatopods of all sizes maximize their flow asymmetry. One consequence of this difference in flow rate is that the water surrounding the aesthetascs at the end of a flick is different from the water surrounding them before the flick, ensuring discrete sampling of odor signals (see also Mead et al., 1999
; Goldman and Koehl, 2001). Animals unable to efficiently clear their aesthetascs of previously sampled fluid are likely to lose temporal and spatial information. |
NEW DIRECTIONS |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
Molecular flux at the aesthetasc surface
The physical model experiments lead to an understanding of the flow patterns around aesthetascs during flicking. Since molecular arrival at the aesthetasc surface involves both advection and molecular diffusion, we created a mathematical model. Velocity field data from the physical model experiments were incorporated into an advection-diffusion solver that calculates molecular flux at the aesthetasc surface (Stacey et al., 2002
). Figure 5A shows one frame in a time series of an odor filament moving through an aesthetasc array. We found that molecular arrival at a sensillar surface was faster during the flick outstroke than during the return stroke, faster for the adult morphology than for the juvenile morphology, and faster at the aesthetasc tip than at the aesthetasc base.
|
Effects of ambient flow on flicking
The aesthetasc Re values that facilitate pulsatile odor sampling rely on the relative velocity of the antennule to the ambient flow. To examine the effects of ambient flow on flicking, I subjected H. ensiguera to environmentally relevant currents in the lab, and then filmed the same species flicking in the field in oscillating flow. Preliminary results suggest that as ambient flow in the direction of the flick outstroke increases, H. ensiguera increase their outward flicking velocity to nearly match the ambient flow but decrease their return stroke flicking velocity. These actions mean that the antennule velocity relative to the surrounding fluid is faster on the return stroke of the flick than on the flick outstroke. As ambient flow in the direction of the return stroke increases, H. ensiguera decrease their outward flicking velocity and increase their return stroke flicking velocity to nearly match the ambient flow. These changes cause the antennule velocity relative to the surrounding fluid to be faster on the flick outstroke than on the return stroke. The changes in antennule velocity with ambient flow ensure that there is always a large asymmetry in relative velocity and thus volume flow rate between the two strokes of a flick (Mead, unpublished data).
Behavioral experiments in an environmental flume
Figure 5B shows a path taken by a H. ensiguera as it navigates up an odor plume in a flume in oscillating flow. Animal orientation, velocity, flicking, and turning as functions of both position in the odor plume and flow characteristics are being analyzed (Mead and Wiley, in preparation). Figure 5C shows a close-up image of H. ensiguera tracking a similar odor plume illuminated by a light sheet. We simultaneously measured odor concentration along the aesthetasc array on the antennule, local instantaneous fluid velocity, and animal behavior (locomotory speed, turning, flicking, etc.). We found that mantis shrimp encounter signal along their antennules differently in wave-affected flow and in unidirectional flow (Mead et al., submitted), and that plume-tracking success varied with flow conditions. We hope that these data will help us understand which cues (such as maximum or average odor concentration along the antennule, filament width, rate of change of odor concentration as a function of space or time, frequency of filament interception, etc.) are required by stomatopods to trace odor plumes.
|
CONCLUSIONS |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
This approach of using techniques from several disciplines to investigate stomatopod olfaction has led to several insights about antennule design and function. For instance, both electron microscopy and fluid measurements were necessary to observe that the aesthetasc cuticle is thinnest and the outer dendritic segments most highly branched at the portion of the aesthetasc with the greatest fluid penetration. In another example, the fact that stomatopods in the field alter their flicking velocity to compensate for rapidly changing ambient flow and thus maintain high volume flow rate ratios between the two strokes of the flick serves to underline the importance of stroke asymmetry derived from the physical model data.
|
ACKNOWLEDGMENTS |
|---|
I would like to thank M. A. R. Koehl for her support and guidance during the years of this research project, and Mark Stacey and Meg Wiley for letting me include data from our joint projects. I also thank the organizers of this symposium for the opportunity to participate. This research was supported by ONR grants N00014-96-1-0594 and N00014-98-1-0775 to M. A. R. Koehl.
|
FOOTNOTES |
|---|
1 From the Symposium Molecules, Muscles, and Macroevolution: Integrative Functional Morphology presented at the Annual Meeting of the Society for Integrative and Comparative Biology, 3–7 January 2001, at Chicago, Illinois.
2 E-mail: kmead{at}socrates.berkeley.edu
|
References |
|---|
|
TOP
SYNOPSISINTRODUCTIONSTOMATOPOD CHEMOSENSOR...FLUID ACCESS TO SENSORSNEW DIRECTIONSCONCLUSIONSReferences |
|---|
Atema, J. 1985. Chemoreception in the sea: Adaptations of chemoreceptors and vehavior to aquatic stimulus conditions. Soc. Exp. Biol. Symp, 39:3887-3423.
Berg, H. C. 1983. Random walks in biology. Princeton University Press, Princeton.
Caldwell, R. L. 1979. Cavity occupation and defensive behaviour in the stomatopod Gonodactylus festae: Evidence for chemically mediated individual recognition. Anim. Behav, 27:194-201.[CrossRef]
Caldwell, R. L. 1985. A test of individual recognition in the stomatopod Gonodactylus festae. Anim. Behav, 33:101-106.
Caldwell, R. L. 1987. Assessment strategies in stomatopods. Bull. Mar. Sci, 41:135-150.
Caldwell, R. L., C. G. Roderick, and S. M. Shuster. 1989. Studies of predation by Gonodactylus bredini. In E. A. Ferrero (ed.), Biology of stomatopods. Collana UZI, Selected Symposia and Monographs pp. 117–131. Mucchi Editore, Modena.
Cheer, A. Y. L., and M. A. R. Koehl. 1987. Paddles and rakes: Fluid flow through bristled appendages of small organisms. J. Theor. Biol, 129:17-39.[CrossRef]
Derby, C. D. 1982. Structure and function of cuticular sensilla of the lobster Homarus americanus. J. Crust. Biol, 2:1-21.[Medline]
Derby, C. D., H. S. Cate, and L. R. Gentilcore. 1997. Perireception in olfaction: Molecular mass sieving by aesthetasc sensillar cuticle determines odorant access to receptor sites in the caribbean spiny lobster Panulirus argus. J. Exp. Biol, 200:2073-2081.[Abstract]
Devine, J. P., and J. R. Koseff. 2001. High resolution measurements of the spatial and temporal scalar structure of a turbulent plume. Exp. In Fluids, 31:90-102.[CrossRef]
Gleeson, R. A. 1982. Morphological and behavioral identification of the sensory structures mediating pheromone reception in the blue crab, Callinectes sapidus. Biol. Bull, 163:162-171.
Gleeson, R. A., W. E. S. Carr, and H. G. Trapido-Rosenthal. 1993. Morphological characteristics facilitating stimulus access and removal in the olfactory organ of the spiny lobster, Panulirus argus: Insight from the design. Chem. Senses, 18:67-75.
Gleeson, R. A., L. M. McDowell, and H. C. Aldrich. 1996. Structure of the aesthetasc (olfactory) sensilla of the blue crab, Callinectes sapidus: Transformations as a function of salinity. Cell Tissue Res, 284:27-288.[CrossRef]
Goldman, J., and M. A. R. Koehl. 2001. Fluid dynamic design of lobster olfactory organs: High-speed kinematic analysis of antennule flicking by Panulirus argus. Chem. Senses, 26:385-398.
Gomez, G., R. Voigt, and J. Atema. 1999. Temporal resolution in olfaction III: Flicker fusion and concentration-dependent synchronization with stimulus pulse trains of antennular chemoreceptor cells in the American lobster. J. Comp. Physiol. A, 185:427-436.[CrossRef]
Grünert, U., and B. W. Ache. 1988. Ultrastructure of the aesthetasc (olfactory) sensilla of the spiny lobster, Panulirus argus. Cell Tissue Res, 251:95-103.
Hallberg, E., K. U. I. Johansson, and R. Elofsson. 1992. The aesthetasc concept: Structural variations of putative olfactory receptor cell complexes in Crustacea. Micros. Res. and Tech, 22:325-335.
Hallberg, E., K. U. I. Johansson, and R. Wallén. 1997. Olfactory sensilla in crustaceans: Morphology, sexual dimorphism, and distribution patterns. Int. J. Insect Morphol. & Embryol, 26:173-180.[CrossRef]
Hansen, B., and P. Tiselius. 1992. Flow through feeding structures of suspension feeding zooplankton: A physical model approach. J. Plankton Res, 14:821-834.
Kaissling, K-E. 1998a. Flux detectors versus concentration detectors: Two types of chemoreceptors. Chem. Senses, 23:99-111.
Kaissling, K-E. 1998b. Pheromone deactivation catalyzed by receptor molecules: A quantitative kinetic model. Chem. Senses, 23:383-395.
Koehl, M. A. R. 1995. Fluid flow through hair-bearing appendages: Feeding, smelling and swimming at low and intermediate Reynolds numbers. In C. P. Ellington, T. J. Pedley (eds.), Biological fluid dynamics, Vol. 49, pp. 157–182. Soc. Exp. Biol. Symp. Company of Biologists Ltd., Cambridge, U.K.
Koehl, M. A. R. 1996. Small-scale fluid dynamics of olfactory antennae. Mar. Fresh Behav. Physiol, 27:127-141.
Laverack, M. S. 1988. The diversity of chemoreceptors. In J. Atema (ed.), Sensory biology of aquatic animals, pp. 287–317. Springer-Verlag, New York,.
Mead, K. S., and M. A. R. Koehl. 2000. Particle image velocimetry measurements of fluid flow through a model array of stomatopod chemosensory sensilla. J. Exp. Biol, 203:3795-3808.[Abstract]
Mead, K. S., M. A. R. Koehl, and M. J. O'Donnell. 1999. Stomatopod sniffing: The scaling of chemosensory sensilla and flicking behavior with body size. J. Exp. Mar. Bio. Ecol, 241:235-261.[CrossRef]
Mead, K. S., and T. M. Weatherby. 2002. The morphology of stomatopod chemosensory sensilla facilitates fluid sampling. Inv. Biol. (In press).
Moore, P. A., J. Atema, and G. A. Gerhardt. 1991. Fluid dynamics and microscale odor movement in the chemosensory appendages of the lobster, Homarus americanus. Chem. Senses, 16:663-674.
Moore, P. A., M. J. Weissburg, J. M. Parrish, R. K. Zimmer-Faust, and G. A. Gerhardt. 1994. Spatial distribution of odors in simulated benthic boundary layer flows. J. Chem. Ecol, 20:255-279.[CrossRef][Web of Science]
Rospars, J.-P., V. K
ivan, and P. Lánsk
. 2000. Perireceptor and receptor events in olfaction. Comparison of concentration and flux detectors: A modeling study. Chem. Senses, 25:293-311.
Schmitt, B. C., and B. W. Ache. 1979. Olfaction: Responses of a decapod crustacean are enhanced by flicking. Science, 205:204-206.
Snow, P. J. 1973. The antennular activities of the hermit crab, Pagarus alaskiensis (Benedict). J. Exp. Biol, 58:745-766.
Sokal, R. R., and F. J. Rohlf. 1995. Biometry, 3rd ed. W. H. Freeman & Company. New York.
Stacey, M. T., K. S. Mead, and M. A. R. Koehl. 2002. Molecular capture by olfactory antennules: Mantis shrimp. J. Math. Biol. 44:1–30.
Vogel, S. 1994. Life in moving fluids, 2nd ed. Princeton University Press, Princeton, N.J.
Weissburg, M. J. 2000. The fluid dynamical context of chemosensory behavior. Biol. Bull, 198:188-202.[Abstract]
Weissburg, M. J., and R. K. Zimmer-Faust. 1993. Life and death in moving fluids: Hydrodynamic effects on chemosensory-mediated predation. Ecology, 74:1428-1443.[CrossRef]
Weissburg, M. J., and R. K. Zimmer-Faust. 1994. Odor plumes and how blue crabs use them in finding prey. J. Exp. Biol, 197:349-375.[Abstract]
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  M.A.R. Koehl The Fluid Mechanics of Arthropod Sniffing in Turbulent Odor Plumes Chem Senses, February 1, 2006; 31(2): 93 - 105. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. A. Keller and M. J. Weissburg Effects of Odor Flux and Pulse Rate on Chemosensory Tracking in Turbulent Odor Plumes by the Blue Crab, Callinectes sapidus Biol. Bull., August 1, 2004; 207(1): 44 - 55. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||