Principles underlying chromatophore addition during maturation in the European cuttlefish, Sepia officinalis

Camouflage plays a key role in the lives of many animals both as a method of defense enabling prey to hide from predators and as a hunting strategy allowing predators to stalk prey without detection (Stevens and Merilaita, 2009). One important aspect of evading detection by predators is the use of body patterning, in which colored patterns are displayed by an organism on its exterior (Cott, 1940). Although there are countless numbers of different camouflage body patterns to help animals hide in their natural environments, often these patterns can be categorized into three basic classifications: uniform, mottle and disruptive (Cott, 1940; Hanlon and Messenger, 1988; Barbosa et al., 2007). The vast majority of animals utilize only one static body pattern expressed throughout their lives regardless of environmental changes (Carvalho et al., 2006; Herbert, 1974; Johnsen, 2001; Sazima et al., 2006; Fox and Vevers, 1960). Some species can slowly change their coloration to reflect changes in the environment, e.g. the snowshoe hare, which alters its fur color from rusty brown in the summer to solid white in the winter (Severaid, 1945). Only a small percentage of animals, however, can actively change their coloration in real-time to reflect variations in their background.

Coleoid cephalopods, unshelled mollusks including cuttlefish, squid and octopus, are among the very few organisms capable of changing their skin color and texture to match their background (Boycott, 1961; Hanlon and Messenger, 1996; Messenger, 2001). The control of body patterning is so rapid – less than 1 s – that these organisms are capable of producing dynamic body patterns such as the ‘passing wave’, which mimics the movement of waves on the ocean floor (Hanlon and Messenger, 1988).

Cephalopods achieve sophisticated control over their body patterning in large part because of the presence in their skin of three different classes of specialized coloration cells known as iridiphores, leucophores and chromatophores (Brocco and Cloney, 1980; Packard, 1985; Hanlon and Messenger, 1988). Of these, the chromatophores are perhaps the most interesting because of their unique ability to rapidly change their shape through a specialized neuromuscular control system. Chromatophore cells have several, unique cellular specializations. Each contains one of three classes of cytoplasmically localized pigment molecules: black, reddish-brown or yellow. Attached to the highly infolded and very elastic plasma membrane are 6–20 radially arrayed chromatophore muscles directly innervated by the central nervous system (Cloney and Florey, 1968). Simultaneous contraction of all chromatophore muscles results in the rapid expansion of the pigmented chromatophore cell up to 10 times its original diameter, from ∼30 μm when condensed to ∼300 μm when fully expanded (Cloney and Florey, 1968; Loi and Tublitz, 1998). A single chromatophore cell, its surrounding chromatophore muscles and adjacent support cells are collectively referred to as a chromatophore organ (Cloney and Florey, 1968).

While all unshelled cephalopods have chromatophore cells and chromatophore organs, their number, distribution and function vary during post-embryonic maturation and across species (Packard and Sanders, 1969; Packard, 1985; Hanlon and Messenger, 1988; Shohet et al., 2007). At birth, a cuttlefish hatchling is generally less than a few centimeters in size and contains only several hundred chromatophores on its entire body (Hanley et al., 1998; Domingues, 2001). As the individual grows, new chromatophores are added until each adult has hundreds of thousands of chromatophores spread across its body in a precise array, with each color class of chromatophore lying in its own epidermal layer (Cloney and Florey, 1968; Packard and Hochberg, 1977; Cloney and Brocco, 1983; Packard, 1985; Hanlon and Messenger, 1996). Regular location and spacing of chromatophores is essential for accurate and functional body patterning. Any gap or disruption in the chromatophore array could substantially compromise the cuttlefish's ability to camouflage properly, increasing the risk of predation and making it difficult to capture prey (Hanlon, 2007). For this reason it is crucial for new chromatophores to be added in a non-random manner to maintain consistent chromatophore spacing necessary to generate seamless body patterns.

The purpose of this study was to determine the general principles underlying the growth and addition of new chromatophore cells. Every new chromatophore is seamlessly inserted and functionally integrated into the existing chromatophore matrix within a few days after appearance without hindering the ability of the organism to produce body patterns. The addition of so many new, functional chromatophores into a pre-existing chromatophore system raises multiple developmental and neural control issues including the precision of their insertion location and the ability of the nervous system to precisely wire up new chromatophores into the existing array. We focused on the former issue as a foundation for a future investigation into the latter. This study used the European cuttlefish, Sepia officinalis, because it is easily reared in the laboratory and because of the wealth of literature on its body coloration patterns. Chromatophore addition was analyzed in normal growth and regeneration because each provided different experimental conditions to explore and understand the principles underlying this interesting phenomenon. The results of this study should enhance our understanding of the mechanisms by which cephalopods achieve adaptive body patterning for camouflage and communication.

Fourteen European cuttlefish, S. officinalis L. (age at beginning of study: mean 3 months, range 2 weeks to 7 months; mean mantle length 7.5 cm; range 3.2–9.1 cm) were used for filming and observation of fin growth. All test subjects were purchased from the National Resource Center for Cephalopods, Marine Biomedical Institute, Galveston, TX, USA, and were housed in two, 473 l artificial seawater (ASW) tanks (122×60×60 cm) at the Institute of Neuroscience at the University of Oregon. Water conditions and culturing practices were identical to those previously described (Loi and Tublitz, 1998). Water temperature was maintained at 20–21°C, and animals were fed a mixture of live zebra fish and frozen shrimp twice a day. Test subjects were held together unless fighting required the insertion of a Plexiglas® wall to separate them. This study was carried out over the course of a year and a half with filming occurring during each of the four seasons. Mantle length was monitored throughout the 30 day filming period and most animals added 1–2 cm to their mantle length during this period.

To analyze chromatophore addition during regeneration, a ∼3 mm square piece of tissue from one swimming fin was excised from each of seven juvenile cuttlefish. This was accomplished by first removing animals from their home tanks and placing them in a glass dish with home tank seawater. Animals were anesthetized by slowly adding 95% ethanol to the seawater. The anesthetic was administered slowly until the animal's breathing pattern, color and response to touch indicated that it was sufficiently sedated for filming. A ∼3 mm square of dermis was removed from the anterior fin using angled iris dissection scissors with the aid of stainless steel forceps (Dumont #5 Dumoxel). After surgery, each animal was transferred to a round, glass dish (65×28 cm diameter×height) with clean tank water and returned immediately to their home tank. Including time to fully anesthetize the animal, the entire surgical procedure took 1–1.5 h to complete.

Re-growth of the fin dermis into the excised region was monitored by filming every 3–4 days for the duration of the regeneration period, ∼28–30 days, with day 0 being the day the fin tissue was removed. Individuals were removed from their home tank and anesthetized following the procedure described above for the fin excision surgery. Video images were taken using a flexcam (VideoLabs, Golden Valley, MN, USA) connected to a digital camera (Sony NP-F750, Tokyo, Japan) on a dissecting microscope (Leica Wild M3C, Wetzlar, Hesse, Germany). Because ethanol anesthesia caused the cuttlefish muscles to relax, video recordings and ICD measurements were taken on fully condensed chromatophores. Lighting and magnification levels were adjusted during each filming to maximize image clarity and visibility of chromatophores. Directed light (Fiberlite, Dollan Jenner, Boxborough, MA, USA) was used when needed. Still images were obtained from video recordings using iMovie (Apple) and Windows Media Player (Microsoft) software.

Seven cuttlefish were filmed three times a week for a period of 30 days using a charge-coupled device (CCD) digital microscope camera (Moticam 2000, Richmond, BC, Canada) centrally positioned within the eyepiece of a dissection microscope (Wild M3C). Animals were individually brought from their 473 l home tank to the filming area in a small seawater-containing dish and were lightly anesthetized using 95% ethanol. Anesthesia was monitored as described in the previous section. Following sedation, the animal was transferred to a shallow, rounded glass filming dish (41×16 cm diameter×height) filled with seawater. The dish contained an oval groove made within a silicone base (Sylgard 182 elastomer) to hold the body of the cuttlefish ventral side down. This position allowed the fin to be raised and flattened across a marked 1×1 mm grid to standardize measurements during filming.

Data were collected from observations on a 2×2 mm section of the posterior fin. This location was chosen because it is an area with very little movement, is easily identifiable, and is far from the animal's eyes and head, making filming under lights less stressful for the test subject. Using bright directed light from a fiberoptic light source (Fiberlite), video was taken at 20 frames s^–1 with a Moticam 2000 CCD camera attached to a Wild M3C dissecting microscope at a magnification of ×400. Three or four adjacent chromatophores were chosen as visual landmarks on the first day of filming for each animal and each subsequent day of filming captured these selected chromatophores in the same frame for comparative purposes. This allowed the same 2×2 mm area of fin to be unequivocally identified during filming. Video from each day's filming was transferred to a computer and individual still images from the video were captured using Windows Media Player and a snipping program. Captured still frames were printed out in color and used for data analysis.

For each day of filming, 100 randomly selected black chromatophores on the same fin as the experimental data but away from the excision/regeneration site were identified and mean interchromatophore distance (ICD) computed. ICD was used as a measure of spatial frequency to quantify the relative distance between each chromatophore and its neighbors. Using the still images obtained from the filming of normally developing or regenerating fin tissue, mean ICD for each black chromatophore was determined by measuring the mean distance in millimeters between its center and the center of each of its three nearest neighboring black chromatophores. Raw measurements were taken using Adobe Photoshop CS. These data were then transferred to Microsoft Excel for data analysis. This same procedure was also used to determine the mean daily control ICD for 100 yellow and 100 reddish-brown chromatophores using the three closest chromatophores of the same color. Separate control data sets were collected for the normal growth and regeneration parts of this study.

For each day of filming, mean ICDs in millimeters were obtained for every new chromatophore when it first appeared in the regenerating piece of fin tissue. One-hundred existing chromatophores of the same color were randomly selected each filming day from regions adjacent to the excision site and used for control data. Selection of adjacent chromatophores was avoided so that all control data were independent. For each of the new and control chromatophores, the distances to its three closest neighboring chromatophores of the same color were recorded and averaged to find the mean ICD. When comparing still photographs from different days, obvious chromatophore clusters and leucophores were used as recognizable markers around the excised region. This process made it easier to identify new chromatophores. Because there was slight variance in the ICD of chromatophores closest to the edge of the fin compared with more interior chromatophores, control chromatophores were randomly selected from all regions of the fin surrounding the cut site to avoid skewed control measurements. All data were statistically analyzed using a one-way ANOVA in Microsoft Excel.

Each new black, reddish-brown and yellow chromatophore in the 2×2 mm filmed region was individually identified and tracked chronologically through to day 30. Every new chromatophore was given a number in the order of its initial appearance in the skin. Nearly all new chromatophores first appeared as a faint orange color not recognizable as any of the three chromatophore colors (see Results and Discussion). After several days, before any substantial growth in chromatophore size had occurred, each new chromatophore became unmistakably recognizable as black, reddish-brown or yellow. ICD measurements for every new chromatophore were obtained using chromatophores of that same color class and collected on the day of its initial appearance only after the color of a new chromatophore was permanently identifiable. All data were statistically analyzed using one-way ANOVA in Microsoft Excel.

Coleoid cephalopods have the very rare ability to completely regenerate their dermis after substantial damage (Sereni and Young, 1932). For example, following excision of a section of the fin from a juvenile S. officinalis, fin re-growth occurs quickly and the new growth becomes fully functional and integrated with surrounding tissues within roughly 30 days (Fig. 1). We used this rapid regeneration ability as a tool to identify a few principles involved in chromatophore addition. Re-growth and placement of chromatophores were monitored following the removal of a small piece of fin tissue from juvenile S. officinalis as described in Materials and methods. This experiment focused exclusively on the addition of black chromatophores. Unequivocal identification of individual new black chromatophores was facilitated by the presence of existing black chromatophores because the latter did not migrate or change location in relation to new chromatophores and because they were easily used, along with stationary leucophores, as recognizable markers around the excised region.

The results of the regeneration experiments demonstrate a significant difference between the mean ICD of control (i.e. existing) and newly added black chromatophores (Figs 2 and 3). Mean ICDs from newly inserted chromatophores were substantially greater than those of controls during the first few days of this study and remained so for the first two-thirds of the observation period (days 0–16). Beginning on day 20, mean ICDs from new chromatophores became smaller than control values and this relationship continued until the fin was fully regenerated (day 30). Over the 30 day observation period, the mean ICD of newly added black chromatophores declined by 44%, starting at an initial value of 340 μm on day 4 and ending with a mean ICD of 190 μm on day 30 (Figs 2 and 3). There was very little variation in the mean daily ICDs of existing chromatophores through the course of the study (±20 μm s.e.m.). A comparison of the mean ICDs from newly inserted and existing black chromatophores revealed a significant difference between the two values for each day of this study (P<0.01 for all days except day 30 where P<0.05).

Several observations during the regeneration study were noted. Regenerated chromatophores appeared only after tissue re-growth and not simultaneously. New tissue growth began almost immediately, however, with new tissue appearing after only 24 h. New chromatophores were visibly identifiable for every recorded day, including the first day of data collection. This observation differs from that of leucophores, which were not fully restored in the excision site tissue by the end of the filming period. Newly inserted chromatophores were initially observed along the growing edge of the excised region, furthest from existing chromatophores (Fig. 1). Only after substantial tissue re-growth had occurred were new chromatophores detected in more interior regions of the excision site (Fig. 1). New chromatophores tended to emerge in groups of approximately 3–5 chromatophores, though single ones did appear in more interior regions of the regenerating piece of fin. It was also observed that the density of black chromatophores in the excision site increased during regeneration until it was first equivalent to and then greater than that of adjacent, undisturbed fin tissue (Figs 2 and 3).

To determine whether the rules for new chromatophore insertion during regeneration also pertained to normal development, ICDs of newly added fin chromatophores were collected during normal growth and maturation and compared with ICDs from existing (control) chromatophores. Data for each chromatophore type (yellow, reddish-brown and black) were obtained from seven juvenile cuttlefish and analyzed separately. No new chromatophore was used as a data point for ICD analysis until its color became unequivocally distinguishable as either yellow, reddish-brown or black.

As a general rule, approximately half of the new chromatophores were inserted along the outside edge of existing chromatophores and the remainder were added to the interior between pre-existing chromatophores. The overall mean ICD of newly inserted black chromatophores was 150±22 μm (s.e.m.) over the 30 day measurement period (Fig. 4A, Fig. 5). In contrast, the overall mean ICD for existing (control) black chromatophores was significantly larger at 190±36 μm (Fig. 4A, Fig. 5). Daily mean ICDs of new or existing chromatophores did not vary significantly during the data collection period.

The same general pattern was observed for the reddish-brown (Fig. 4B, Fig. 5) and yellow chromatophores (Fig. 4C, Fig. 5). New reddish-brown chromatophores were added at a mean ICD of 190±34 μm compared with their mean control ICD of 210±12 μm, and new yellow chromatophores appeared at a mean ICD of 110±23 μm compared with their control data of 150±46 μm (Fig. 5). Statistical analysis using a one-way ANOVA showed that existing (control) black, yellow and reddish-brown chromatophore ICDs were significantly different from each other (P<0.01). ICDs of control and new chromatophores were also significantly different from each other for each chromatophore color type (Fig. 5), with the ICDs of existing chromatophores being greater than those of their respective newly added chromatophores. The ratios of the mean ICDs of new and existing chromatophores for each color type revealed that new chromatophores were always inserted closer to their nearest neighbors than the normal chromatophore spatial frequency (0.74, 0.85 and 0.57 for black, reddish-brown and yellow chromatophores, respectively). These data also indicate a higher density (i.e. smaller ICD) of yellow chromatophores in the fin than of either black or reddish brown chromatophores (Fig. 4C, Fig. 5).

To understand the maturation process of newly formed chromatophores, we individually tracked 516 chromatophores from their initial appearance to their adult phenotype. New chromatophores always appeared first as very small, pale orange cells, and were not classifiable as any of the three mature chromatophore colors (Fig. 6). Following their initial appearance, all new chromatophores slowly darkened and, in the case of black chromatophores, grew larger over the course of 27–30 days until they were the same color and size as existing chromatophores of the same color type. For example, black chromatophores turned from an initial pale orange to a reddish-orange color, then to dark brown and finally to their mature dark brown–black color, similar to that of surrounding black chromatophores (Fig. 7). This darkening developed slowly over the course of a few weeks rather than in a step-wise manner in which color changed dramatically from day to day. During the time they were darkening, developing black chromatophores were never recognizable as either yellow or reddish-brown chromatophores. New reddish-brown and yellow chromatophores developed similarly to black chromatophores, starting out as small pale orange cells that changed color until they closely resembled surrounding chromatophores of the same color type. Once a chromatophore became recognizable as a fully grown black, reddish-brown or yellow chromatophore, its color remained unaltered for the remainder of the observation period.

To quantify the changes in size as chromatophores matured, the diameters of new chromatophores were measured over time as they developed. The diameters of existing yellow, reddish-brown and black chromatophores were also obtained as controls. Because the ethanol anesthesia procedure causes relaxation of the peripheral musculature including the chromatophore muscles, all chromatophore diameter measurements were performed on completely condensed chromatophores. Newly inserted black chromatophores grew substantially as they developed and matured, enlarging from an initial mean diameter of 17±0.39 μm on day 1 to a diameter of 36±0.57 μm on day 29 (Fig. 8A). Unlike their black counterparts, new reddish-brown and yellow chromatophores did not significantly change in size during maturation, with their diameters remaining statistically similar to those of existing chromatophores throughout the study (diameters of 18±0.32 and 18±0.44 μm, respectively; Fig. 8B,C). On the first day of appearance, the diameters of new reddish-brown, yellow and black chromatophores were all statistically similar to each other (P>0.05).

The objective of this study was to identify a few of the principles underlying chromatophore organization during normal growth. This is a necessary first step towards understanding how the central nervous system mediates body patterning and how chromatophores are organized and arranged in the skin of cephalopods for the twin purposes of camouflage and communication. Chromatophores, when appropriately distributed across the body, are the primary components of the numerous, intricate, body patterns that mediate intraspecies and interspecies communication, including camouflage, mating and prey capture (Hanlon and Messenger, 1988; Boal et al., 2004). In contrast, an irregular or patchy arrangement of chromatophore cells during growth would interfere with the proper production of body patterns, and negatively impact the survival and reproductive success of these organisms. The results of this study strongly support the hypothesis that the chromatic component of body patterning in the European cuttlefish, S. officinalis, is highly organized and that the organizational aspects of chromatophore growth (i.e. density, positioning) are precisely regulated.

Two observations from the fin regeneration study suggest that chromatophore addition may be more actively controlled during dermal regeneration than other dermal elements such as leucophores and iridiphores. First, new chromatophores in regenerating tissue appeared shortly after general tissue growth occurred, in less than a week (Figs 1–3). In contrast, new leucophores and iridophores did not appear until the end of the 30 day study and were not fully regenerated once all chromatophores had re-grown and the study had been completed. This observation suggests the mechanism(s) responsible for chromatophore growth and subsequent innervation during regeneration is either continuously active or activated in immediate response to tissue damage.

The second indication of active regulation was the progressive decrease in the mean experimental ICD during tissue regeneration until it was equal to and then less than that of the surrounding tissue (Figs 2 and 3). The first sets of new chromatophores were inserted in the interior of the regeneration site, relatively far from their neighbors. Subsequent new chromatophores filled in the gaps between the first set of chromatophores until the ICD of new chromatophores in the regeneration site was less than that in control tissue. This observation was similar to that of the normal growth study in which new chromatophores were added at ICDs less than those in controls. This suggests that at this point in regeneration the tissue was acting similarly to normally growing tissue. The acute reduction in the ICD of new chromatophores during regeneration suggests that more than one mechanism may be involved in determining the location of a new chromatophore. If only one primary factor was responsible for new chromatophore positioning, the ICDs of new chromatophores would be the same as control ICDs throughout the entire regeneration process. In such a model, new chromatophores would be added linearly only along the growing edge of the regeneration site close to existing chromatophores. Instead, by adding chromatophores to the interior of the regeneration site at large ICDs and then filling in the gaps, the regenerating site becomes functional sooner than would have been possible in the single mechanism model. To facilitate this more efficient regeneration strategy, it is suspected that the positioning of new chromatophores is controlled by both positive and negative regulators. The presence of positive regulators promotes chromatophore growth and innervation, while negative regulators inhibit chromatophores from developing too close to other chromatophores, thus maintaining a relatively consistent average ICD. The predictable manner in which new chromatophores are inserted relative to existing chromatophores indicates a high level of organization and strong positive evidence that the presence of old, neighboring chromatophores helps determine the location of new chromatophores. This top-down model is consistent with the evidence presented here; however, it must be noted that other models, including a simple attraction–repulsion model, are also possible.

In normally growing tissue, most new chromatophores were added in between existing chromatophores. Some new chromatophores, however, were also inserted along the outside edge of the fin to complete lines and fill in gaps as the fin enlarged. New chromatophores inserted in this manner rarely appeared far away from existing chromatophores and had ICDs close to or less than controls (J.Y., personal observation). Consistent with the results of the regeneration study, this observation provides evidence that the proximity of existing chromatophores is one of the primary effectors regulating the location of new chromatophores. The addition of new chromatophores at smaller than normal ICDs presumably provides for growth without reducing the grain density of chromatophores in body patterns.

The details of chromatophore development in cephalopods remain unresolved, in large part because of a paucity of studies. Two competing hypotheses have been proposed. One study in Octopus vulgaris proposed that all new chromatophores begin as yellow chromatophores, and then turn reddish-brown before finally becoming black chromatophores (Packard, 1985). An alternative hypothesis is that each chromatophore is born as a specific color type and remains that type, e.g. yellow, reddish-brown or black, for its entire lifetime. The data presented in this paper refute the first hypothesis and provide support for the second (Fig. 9).

The majority of new chromatophores observed in this study started out as small, pale orange cells that developed directly into yellow, reddish-brown or black chromatophores without passing through either of the other two colors during the month-long observation period. We looked very closely but did not observe any reddish-brown chromatophores passing through a transient yellow phase or a mature black chromatophore which began yellow in color and went through a reddish-brown phase before becoming black. All yellow chromatophores observed on the first day of filming remained yellow at the end of the 30 day observation period, during which we observed the appearance of many new black chromatophores. The observation that yellow chromatophores did not darken while many new orange-colored chromatophores became fully mature black chromatophores is strong evidence against the idea that groups of chromatophores develop together in waves and all mature together at the same rate while passing through a yellow, reddish-brown and black developmental sequence as proposed by Packard (Packard, 1985). Our data support Packard's observation that new chromatophores are always small and orange-colored (Packard, 1985).

An important observation was that those new chromatophores fated to become black chromatophores doubled their diameters over the course of a few weeks (Fig. 8) while the yellow and reddish-brown chromatophores remained the same size as at their initial appearance. The larger size of the black chromatophores may be due to the need to generate large areas of black on the skin for striped patterns such as the zebra and passing cloud displays (Packard and Sanders, 1971; Hanlon and Messenger, 1996; Hanlon and Messenger, 1988). Each chromatophore color type lies in its own dermal layer, with the black chromatophores being the most superficial followed by the reddish-brown and yellow chromatophore layers (Cloney and Florey, 1968; Cloney and Brocco, 1983; Hanlon and Messenger, 1996; Sutherland et al., 2008). This study was unable to track any vertical movement of chromatophores as they matured. However, given the knowledge that chromatophores of different colors reside in separate dermal layers and the hypothesis proposed by Packard (Packard, 1985) that chromatophores pass through all three colors, this would be an interesting topic for future investigations.

Our data show that chromatophore growth and development are highly organized and structured. From our observations, we propose five guiding principles of chromatophore growth and maturation. (1) Each of the three chromatophore cell types – black, reddish brown and yellow – are present at different spatial frequencies in the cuttlefish fin. (2) During normal growth, new chromatophores are inserted at a higher spatial frequency than existing (control) chromatophores of the same color type. (3) In regenerating tissue, new black chromatophores are initially added at low spatial frequencies. As regeneration continues, new black chromatophores appear at increasing spatial frequencies until they are inserted at a spatial frequency higher than observed in control tissue. (4) All chromatophores first appear as pale orange cells and slowly darken into their respective color types without passing through intermediate color stages. (5) New black chromatophores undergo a doubling in size as they mature while, reddish-brown and yellow chromatophores do not grow at all after they are inserted in the dermis.

Several of these principles have been observed in other studies (Packard, 1985; Hanlon and Messenger, 1988; Messenger, 2001; Domingues, 2001); however, this is the first study to statistically analyze the growth and spatial organization of chromatophores as they develope. The use of modern data capture and analyses techniques allowed a vastly larger number of chromatophores to be tracked and analyzed than was possible in earlier studies. A well-developed understanding of the rules controlling chromatophore addition and positioning is an important first step in elucidating the basic neural organization and processes that underlie the amazingly high level of behavioral plasticity in cephalopods. Although this study did not analyze new chromatophore motoneurons or their innervation pattern, the principles in this study will be a helpful starting point for future examination of this issue. By using the principles outlined in this study it should be possible to make accurate statistical predictions of the location of new chromatophores on the fin. This information should make it easier to predict the location of new neurons and innervations and observe and analyze them as they grow.

Further elucidation of the underlying molecular mechanisms controlling chromatophore insertion and spacing is required to obtain a full understanding of how chromatophore growth takes place and how new chromatophores are integrated within the central nervous system. Many questions still remain about how chromatophores develop and mature. Do all three color cells begin as a pluripotent cell that responds to external factors to differentiate into one of the three colors, or are there three separate cell types that lead to each color? Do new chromatophore cells divide close to the site at which they are needed, or do they migrate from other parts of the body? Because many cephalopod species have varying densities of chromatophores (Cloney and Florey, 1968; Packard and Hochberg, 1977; Cloney and Brocco, 1983; Hanlon and Messenger, 1996; Sutherland et al., 2008), it would also be interesting to see whether the patterns and principles observed in S. officinalis in this study are conserved across other cephalopod species.