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doi:10.2204/iodp.proc.338.102.2014

Biostratigraphy

Because there were no micropaleontologists on board during Expedition 338, micropaleontological investigation occurred on shore after Expedition 338. Calcareous nannofossils and radiolarians in core and cuttings samples were collected from holes at Site C0002 and calcareous nannofossils and planktonic foraminifers in core samples were collected from Holes C0021B and C0022B.

Calcareous nannofossils

During Expedition 338, calcareous nannofossils were used to date the core catcher and cuttings samples. When necessary, we took additional samples from the cores to further refine ages.

Zonation and biohorizons

For Expedition 338, we applied a biostratigraphic zonation of calcareous nannofossils based on the zonal schemes of Martini (1971) and Okada and Bukry (1980). Our application of zonal markers and additional datums is mostly based on the compilation by Raffi et al. (2006) and Raffi (2002), in line with previous Expeditions 315, 316, 319, 322, and 333 for biostratigraphic consistency and subsequent correlation (see “Biostratigraphy” in Expedition 315 Scientists [2009a], Expedition 316 Scientists [2009a], Expedition 319 Scientists [2010b], Expedition 322 Scientists [2010a], and Expeditions 333 Scientists [2012a]).

Each nannofossil datum was assigned an astronomically calibrated age compiled by Raffi et al. (2006). The astrochronological frame for the Neogene follows the International Commission on Stratigraphy 2004 timescale (Lourens et al., 2004). The timescale and biostratigraphic zones of calcareous nannofossils are summarized in Figure F15 and Table T11.

Downhole contamination is common in riser drilling cuttings and often poses problems in recognition of a zonal boundary defined by a first occurrence (FO) datum because such a boundary may appear significantly stratigraphically lower than in situ. To circumvent this problem, a last occurrence (LO) datum stratigraphically close to a FO datum, if available, was selected to approximate the zonal boundary; otherwise, the biozone was combined with adjacent zones.

We applied two additional criteria to resolve the reworking of zonal markers, which tends to make the assemblage appear older. First, a datum was defined by the continuous occurrence of a taxon, whereas sparse occurrence was considered reworked. In addition, we used changes in assemblage composition and specimen size to evaluate the occurrence of zonal markers.

Taxonomic remarks

Taxonomy followed the compilation of Perch-Nielsen (1985) and Young (1998). Previous work suggested grouping reticulofenestrid species including genera Gephyrocapsa and Reticulofenestra by size. This is because their species show a great variation in size and other morphological features (e.g., relative size of the central opening and orientation of the bar in case of Gephyrocapsa).

Accordingly, Gephyrocapsa is divided into three major groups by maximum coccolith length following biometric subdivision by Rio (1982), Raffi et al. (1993) and Raffi (2002): small Gephyrocapsa (<4 µm), medium Gephyrocapsa (4–5.5 µm), and large Gephyrocapsa spp. (>5.5 µm). Some important morphologic features (e.g., bar orientation) were also considered during the analysis. In addition, Reticulofenestra pseudoumbilicus should have a coccolith length >7 µm. Other smaller Reticulofenestra specimens (<7 µm) are included in Reticulofenestra spp., although Reticulofenestra asanoi, a characteristic circular taxon >6 µm in diameter, is separated.

Methods

We prepared smear slides from cuttings sampled at 50 m spacing and core catcher samples within the cored interval, following the standard method with photo-curing adhesive as a mounting medium. A simple concentration technique was adopted from Expedition 316 (see “Biostratigraphy” in Expedition 316 Scientists [2009a]) for samples that contained coarse materials or few to rare nannofossils. This technique involves suspending and settling sediment in distilled water to remove sand and silt before making smear slides.

In addition, we used a “mixing” technique for cuttings samples to better represent the 50 m sampling interval. We soaked a few grams of clean shale chips or drilling mud sediment mixture (when no hard rock pieces were present) in water and stirred to disaggregate and suspend the sample materials. The sample material was ground with a mortar and pestle when needed. The suspension was allowed to settle for 60 s, and then we drew ~3 mL of the suspension using a transfer pipet for making standard smear slides.

We examined slides using an Olympus BX 50 microscope under cross-polarized light and transmitted light at 1500× magnification.

We estimated relative abundances of nannofossil assemblages based on observations in a traverse at 1500× magnification, although usually more than two transverses (more than ~300 fields of view [FOVs]) were browsed for zonal markers and rare species. A letter code was given to each abundance category and defined as follows:

• V = very abundant (>10 specimens per FOV).
• A = abundant (1–10 specimens per FOV).
• C = common (1 specimen per 2–10 FOVs).
• F = few (1 specimen per 11–100 FOVs).
• R = rare (1 specimen per 101–300 FOVs).
• B = barren (no nannofossils per >300 FOVs; for assemblage abundance only).

We only focus on the presence of nannofossil species in each sample and relative abundance of individual species/genus are not examined. The average preservation state of the nannofossil assemblage in each sample was qualitatively categorized and defined as follows:

• VG = very good (no evidence of dissolution and/or overgrowth; no alteration of diagnostic characteristics; all specimens identifiable at the species level).
• G = good (little or no evidence of dissolution and/or overgrowth; only slight alteration of diagnostic characteristics; most specimens [~95%] identifiable at the species level).
• M = moderate (evident etching and/or overgrowth; diagnostic characteristics sometimes altered; broken specimens frequent and delicate forms decreased; however, the majority of specimens identifiable at the species level).
• P = poor (severe dissolution, fragmentation and/ or overgrowth; diagnostic characteristics largely destroyed; many specimens [>50%] not identifiable at the species and/or generic level).

Radiolarians

Radiolarian zones are given in the Neogene North Pacific zonation of Kamikuri et al. (2004, 2007) and in the Cenozoic low-latitude zonation of Sanfilippo and Nigrini (1998) wherever possible. They also provide numerical ages of bioevents, which are tuned to the geomagnetic polarity timescale (GPTS) of Cande and Kent (1995).

Methods

Sample preparation for microscopic examination followed the standard techniques described by Sanfilippo et al. (1985). Samples were treated with hydrogen peroxide (20% H₂O₂) and sodium pyrophosphate (5% Na₄P₂O₇) and heated to boiling. Hydrochloric acid (HCl) may be added to dissolve calcareous components. Disaggregated particles were wet-sieved through a 63 µm mesh. Remaining residues were removed and dried. Undisaggregated sediment was treated again. The clean particles were spread on glass slides and mounted with Entellan-New. Slides were examined with a transmitted light microscope at 100× to 400× magnification. The first 200 specimens encountered in one slide were counted, after which slides were scanned to determine whether other taxa were present.

Total radiolarian abundance in a slide was based on the following categories:

• A = abundant (>500 specimens in a slide).
• C = common (100–500 specimens in a slide).
• R = rare (10–99 specimens in a slide).
• VR = very rare (1–9 specimens in a slide).

Preservation of the radiolarian assemblage was based on the following categories:

• G = good (radiolarians show no sign of dissolution with only minor fragmentation).
• M = moderate (radiolarians show evidence of moderate dissolution with obvious fragmentation).
• P = poor (radiolarians show signs of a high degree of dissolution with very few intact specimens).

Planktonic foraminifers

The planktonic foraminifer zonation of Blow (1969) and astronomically calibrated biohorizons of Neogene planktonic foraminifers compiled by Wade et al. (2011) were applied for this expedition. In addition, useful biohorizons were employed from literature in the field and converted in age to the current GPTS (ATNTS2012; Hilgen et al., 2012).

The LO of pink-colored Globigerinoides ruber was located at 0.12 Ma in the Indian and Pacific Oceans by Thompson et al. (1979) and confirmed by others (i.e., Li et al. [2005] at ODP Site 1143 in the South China Sea). The LOs of Neogloboquadrina asanoi and Globoquadrina dehiscens and the FO of Globoconella inflata modern form were correlated with geomagnetic polarities at ODP Sites 1150 and 1151 off northeast Japan (Motoyama et al., 2004). The coiling direction change of Pulleniatina spp. from sinistral to dextral has been reported just above Chron C2n (Olduvai) in the Boso Peninsula of central Honshu, Japan (Oda, 1977). The first consistent occurrence of Neogloboquadrina acostaensis was compiled by Berggren et al. (1995) and converted in age to the current GPTS. These biozones and biohorizons are shown in Table T12 and Figure F15.

Methods

About 10 cm³ of sediment from core catcher sections was collected for foraminifer analyses. Soft sediment samples were disaggregated using hydrogen peroxide solution and naphtha. Indurated mudstone samples were treated by the sodium tetraphenylborate method (Hanken, 1979). After samples were macerated, each sample was wet-sieved through a screen (63 µm opening). Planktonic foraminiferal specimens >125 µm were observed using a binocular microscope from the dried residues. A total of 100–5000 specimens were observed in each sample.

The relative abundance of planktonic foraminifers in each sample is based on the following categories:

• VA = very abundant (foraminiferal tests are exclusively dominant of sand-size residues).
• A = abundant (foraminiferal tests occupy 10%–50% of sand-size residues).
• C = common (>1000 specimens are observed in the sample).
• R = rare (100–1000 specimens are observed in the sample).
• VR = very rare (<100 specimens in the sample).

Preservation of each sample was recorded by the following criteria:

• P = poor (dissolution of surface structure and fragmentation are observed; most individuals cannot be identified at the species level).
• M = moderate (dissolution and fragmentation are commonly evident; some individuals are hard to identify).
• G = good (no dissolution; fragmentation of individuals has slightly occurred).

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